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U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID) Fort Detrick, Maryland Occupational Health Manual for Laboratory Exposures to Select (BSL-3 & BSL-4) and Other Biological Agents (Part I) 18 April 2011 0701016F.doc 1 Occupational Health Manual for Laboratory Exposures to Select (BSL-3 & BSL-4) and Other Biological Agents Second Edition April 18, 2011 Lead Editor Janice Rusnak, M.D. FIDSA (Lt. Col, USAF (ret)) Clinical Research Management, Inc. Special Immunizations Program, USAMRIID Fort Detrick, MD Contributing Reviewers Ellen Boudreau, M.D. Chief, Special Immunizations Clinic USAMRIID Fort Detrick, MD Mark Goldberg, M.D. Special Immunizations Clinic, USAMRIID Fort Detrick, MD Roger G. McIntosh, M.D., M.O.H. Chief, Competant Medical Authority Medical Support Biosurety, USAMRIID Fort Detrick, MD Phillip Pittman, M.D., M.P.H., (COL, USA (ret)) Clinical Research Unit, USAMRIID Fort Detrick, MD Catherine Wilhelmsen, DVM, PhD Microbiologist/Biosafety Officer Safety, Radiation & Environmental Division, USAMRIID Fort Detrick, MD 0701016F.doc ii Mark Wolcott, PhD Diagnostic Services Division, USAMRIID Fort Detrick, MD Comments and suggestions are appreciated and should be addressed to: Attn: Dr. Janice Rusnak Special Immunizations Program U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID) 1425 Porter Street Fort Detrick, MD 21702-5011 0701016F.doc iii External Reviewers Cheryl S. Barbanel, M.D., MBA, MPH, FACOEM, FACP Associate Professor of Medicine, BUSM Director of Occupational Health Programs, Boston University Director of Boston University Occupational Health Center, Charles River Campus Boston, MA L. Casey Chosewood, M.D. Senior Medical Officer for WorkLife National Institute for Occupational Safety and Health Centers for Disease Control and Prevention Atlanta, GA Col. George W. Christopher, M.D., USAF, MC Special Assistant for Biological Defense Countering WMD Defense Policy Washington, DC M. Patrica Joyce, M.D., FIDSA Medical Officer NCHHSTP/DHAP/HICSB Centers for Disease Control and Prevention Atlanta, GA Anne Varljen, C.R.N., C.I.C. Infection Control Austin, TX 0701016F.doc iv PREFACE The Occupational Health Manual for Laboratory Exposures to Select (BSL-3 & BSL-4) and Other Biological Agents is intended to serve as a reference for physicians in the Special Immunizations Program (SIP) and the U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID) and other health care providers who are called upon to evaluate USAMRIID employees who may have been exposed to biologic agents, toxins, and other infectious pathogens in the laboratory. The manual represents the collective knowledge and experiences of physicians who have worked and are working in the SIP clinic. It has also benefited immensely from the experienced research scientists (in many instances national or international authorities on the specific pathogen) at the Institute. Critical to the appropriate evaluation and management of a laboratory exposure is an understanding that the process involves a coordinated team effort including the patient, the attending physician, the patient’s work supervisor, the principal investigator (the responsible scientist for the agent of exposure), the safety specialists, and, as appropriate, community health authorities. In some cases, a USAMRIID subject matter expert is involved in an assessment of the circumstances surrounding the exposure and its consequences for the benefit of the patient, the health care team, the Institute, and the local community. The inevitable evolution in technology and medicine will make it necessary to continuously update the contents of this manual. Although every effort will be made to keep the manual current; health care providers are encouraged to use this manual as a guide and to obtain the latest information available through supplemental sources so that they can deliver the highest quality care to those seeking assistance or treatment for exposures to a biological agents being studied at USAMRIID. 0701016F.doc v ACKNOWLEDGEMENTS This manual has been made possible with the generous assistance and support of David DeShazer, Ph.D., Thomas Geisbert, Ph.D., Henry Heine, Ph.D., Marty Heiberg (Center for Infectious Disease Research and Policy), John Hewetson, Ph.D., George Ludwig, Ph.D., Joseph Mangiafico, Mark Poli, Ph.D., Herbert Thompson, Ph.D., Dave Waag, Ph.D., Robert Wannemacher, Ph.D., Catherine Wilhelmsen, DVM, Ph.D., (Microbiologist/Biosafety Officer; Safety, Radiation & Environmental Division), Mark Wolcott, Ph.D. (Diagnostic Services Division), and many others. The exclusion of anyone is purely accidental and in no way lessens the gratitude we feel for contributions received. DISCLAIMER Although every effort has been made to make the information in this manual consistent with official policy and doctrine, the information contained in this manual is not official policy or doctrine of the Department of the Army nor should it be construed as such. As you review this manual, you will find specific therapies and prophylactic regimens for the diseases mentioned. The majority of these are based on standard treatment guidelines; however, some of the regimens may vary from information found in standard reference materials. The reason for this difference is that the clinical presentation of certain diseases resulting from a laboratory exposure may vary from the endemic form of the disease. For ethical reasons, human challenge studies can only be done with a limited number of these agents. Therefore, treatment and prophylactic regimens may be derived from in vitro data, animal models, and limited human data. Occasionally you will find various investigational new drug products mentioned. They are often used in the laboratory setting to provide potential protection to laboratory workers and health care providers. These products are not available commercially and can be given only under a specific protocol with informed consent. They are mentioned for scientific completeness of the manual and are not necessarily to be construed as recommendations for therapy. The guidelines recommended for therapies and dosages may change based on individual circumstances of the patient being treated, site of infection, organism resistance, or development of new drugs and treatment regimens. If possible, consultation with an infectious disease physician is highly recommended before treatment is administered. 0701016F.doc vi Table of Contents Table of Contents Preface .......................................................................................................................................................... v Acknowledgements ...................................................................................................................................... vi Disclaimer .................................................................................................................................................... vi Abbreviations and Definition of Terms ........................................................................................................ x O VERVIEW OF M ANUAL AND L ABORATORY-ACQUIRED E XPOSURES AND I NFECTIONS AT F ORT DETRICK ............................................................................ 1-1 Purpose of the Manual ......................................................................................................................... 1-1 Background of the SIP ........................................................................................................................ 1-1 Summary of Laboratory Exposures and Illnesses at Fort Detrick ....................................................... 1-1 Protection from Laboratory Exposures and Prevention of Illness at USAMRIID .............................. 1-3 Comprehensive Safety Program ................................................................................................... 1-3 SIP-Managed Vaccination and Post-Exposure Prophylaxis Program .......................................... 1-4 User’s Guide to Manual....................................................................................................................... 1-6 Name of Agent .............................................................................................................................. 1-6 References ..................................................................................................................................... 1-6 R ISK ASSESSMENT OF L ABORATORY E XPOSURES AT USAMRIID ......................................... 2-1 Development of Assessment Guidelines.................................................................................................... 2-1 Evaluation of a Potential Laboratory Exposure or Illness at USAMRIID ................................................. 2-1 Stepwise Evaluation of a Potential Laboratory Exposure .......................................................................... 2-1 Step 1: Initial Evaluation .................................................................................................................... 2-2 Step 2: Assessment of Risk ................................................................................................................ 2-2 Assessment of Risk of Exposure ............................................................................................ 2-3 Assessment of Risk of Disease from Infectious Agents ......................................................... 2-5 Assessment for Post-Exposure Prophylaxis ........................................................................... 2-5 Step 3: Medical Management ............................................................................................................. 2-6 Step 4: Corrective Actions.................................................................................................................. 2-7 Step 5: Maintenance of Records Documenting Potential Laboratory Exposure Event ...................... 2-7 Management of Individuals with Febrile Illness That May Be the Result of a Laboratory Exposure .............................................................................................................................................. 2-7 Isolation .................................................................................................................................. 2-7 Initial Evaluation, Assessment, and Laboratory Tests ........................................................... 2-7 Treatment ............................................................................................................................... 2-7 References .............................................................................................................................. 2-7 M ANAGEMENT OF L ABORATORY E XPOSURES AND I LLNESSES FROM BACTERIAL AND R ICKETTSIAL AGENTS ............................................................... 3-1 Anthrax—Bacillus anthracis ............................................................................................................... 3-1 Overview of the Organism and Laboratory Exposures ................................................................. 3-1 The Disease ................................................................................................................................... 3-3 Management of Potential Exposures to B. anthracis (Asymptomatic Persons) ........................... 3-7 Management of Suspected Anthrax Disease (Symptomatic Persons) .......................................... 3-9 Environmental Decontamination and Infection Control ............................................................... 3-9 References ................................................................................................................................... 3-11 Plague—Yersinia pestis ..................................................................................................................... 3-16 Overview of the Organism and Laboratory Exposures ............................................................... 3-16 The Disease ................................................................................................................................. 3-17 Management of Potential Exposure to Y. pestis (Asymptomatic Persons) ................................. 3-19 Management of Suspected Plague (Symptomatic Persons) ........................................................ 3-20 0701016F.doc vii Table of Contents Environmental Decontamination and Infection Control ............................................................. 3-22 References ................................................................................................................................... 3-22 Tularemia—Francisella tularenis ..................................................................................................... 3-26 Overview of the Organism and Laboratory Exposures ............................................................... 3-26 The Disease ................................................................................................................................. 3-29 Management of Potential Exposure to F. tularensis (Asymptomatic Persons) .......................... 3-33 Management of Suspected Tularemia Disease (Symptomatic Persons) ..................................... 3-34 Environmental Decontamination and Infection Control ............................................................. 3-36 References ................................................................................................................................... 3-36 Glanders—Burkholderia mallei ........................................................................................................ 3-40 Overview of the Organism and Laboratory Exposures ............................................................... 3-40 The Disease ................................................................................................................................. 3-41 Management of Potential Exposure to B. mallei (Asymptomatic Persons) ................................ 3-43 Management of Suspected Glanders (Symptomatic Persons) .................................................... 3-45 Environmental Decontamination and Infection Control ............................................................. 3-48 References ................................................................................................................................... 3-48 Melioidosis—Burkholderia pseudomallei......................................................................................... 3-52 Overview of the Organism and Laboratory Exposures ............................................................... 3-52 The Disease ................................................................................................................................. 3-53 Management of Potential Exposure to B. pseudomallei (Asymptomatic Persons) ..................... 3-56 Management of Suspected Melioidosis (Symptomatic Persons) ................................................ 3-58 Environmental Decontamination and Infection Control ............................................................. 3-61 References ................................................................................................................................... 3-61 Brucellosis—Brucella ....................................................................................................................... 3-67 Overview of the Organism and Laboratory Exposures ............................................................... 3-67 The Disease ................................................................................................................................. 3-69 Management of Potential Exposures to Brucella (Asymptomatic Persons) ............................... 3-70 Management of Suspected Brucellosis (Symptomatic Persons) ................................................. 3-71 Environmental Decontamination and Infection Control ............................................................. 3-72 References ................................................................................................................................... 3-73 Q Fever—Coxiella burnetii ............................................................................................................... 3-76 Overview of Laboratory Exposures ............................................................................................ 3-76 The Disease ................................................................................................................................. 3-78 Management of Potential Exposure to C. burnetii (Asymptomatic Persons) ............................. 3-80 Management of Suspected Q Fever Disease (Symptomatic Persons) ........................................ 3-82 Environmental Decontamination and Infection Control ............................................................. 3-84 References ................................................................................................................................... 3-84 APPENDICES Appendix A – Laboratory Exposure Reporting Requirements ................................................................. A-1 Appendix B – General Sample Collection ................................................................................................ B-1 Appendix C – Containment Care .............................................................................................................. C-1 0701016F.doc viii Table of Contents List of Tables Table 1-1. Table 2-1. Table 2-2. Table 2-3. Table 2-4. Table 3-1. Table 3-2. Table 3-3. Table 3-4. Table 3-5. Table 3-6. Table 3-7. Table 3-8. Table 3-9. Table 3-10. Table 3-11. Table 3-12. Table 3-13. Table 3-14. Table 3-15. Table 3-16. SIP Vaccines and Post-vaccination Clearance Levels before Laboratory Entry................. 1-5 Documentation of a Detailed History of the Laboratory Exposure Event .......................... 2-3 Risk Stratification of Percutaneous, Cutaneous, and Mucosal Exposures to Infectious Agents ................................................................................................................ 2-4 Risk Stratification of Aerosolized Exposures to Infectious Agents .................................... 2-4 Assessment of Factors Influencing Risk of Disease after Exposure to an Agent ............... 2-5 Clinical Features of the Major Forms of Anthrax ............................................................... 3-4 Recommended Chemoprophylaxis of Aerosol Exposure to B. anthracis at USAMRIID .. 3-8 Diagnostic Testing for Anthrax ......................................................................................... 3-14 Clinical Features of the Major Forms of Plague ............................................................... 3-18 Diagnostic Testing for Plague ........................................................................................... 3-24 Clinical Features of the Major Forms of Tularemia .......................................................... 3-31 Diagnostic Testing for Tularemia ..................................................................................... 3-38 Clinical Features of the Major Forms of Glanders ............................................................ 3-41 Chemoprophylaxis for Post-exposure to B. mallei............................................................ 3-44 Diagnostic Testing for Glanders ....................................................................................... 3-50 Clinical Features of the Major Forms of Melioidosis ....................................................... 3-54 Diagnostic Testing for Melioidosis ................................................................................... 3-64 Clinical Features of Brucellosis ........................................................................................ 3-69 Diagnostic Testing for Brucellosis .................................................................................... 3-75 Clinical Features of Q Fever ............................................................................................. 3-79 Diagnostic Testing for Q Fever ......................................................................................... 3-88 List of Figures Figure 1-1. Figure 2-1. Figure 2-2. Figure 3-1. Figure 3-2. Figure 3-3. Figure 3-4. Figure 3-5. 0701016F.doc FDA-Licensed and IND Products for Pre- and Post-exposure Prophylaxis........................ 1-5 Stepwise Evaluation of Potential Laboratory Exposure...................................................... 2-1 Decision Tree for Recommending Post-exposure Antibiotic Prophylaxis in Potential Exposures to Bacterial and Rickettsial Agents ................................................................... 2-6 Licensed Anthrax Vaccine for Pre-exposure Prophylaxis .................................................. 3-3 Anthrax Day 3 Skin Lesion ................................................................................................. 3-5 IND Tularemia Vaccine for Pre-exposure Prophylaxis .................................................... 3-27 Day 7 Reaction to Tularemia Vaccination ........................................................................ 3-28 Positive Q Fever Skin Test................................................................................................ 3-78 ix Abbreviations and Definition of Terms ABBREVIATIONS AND DEFINITION OF TERMS Abbreviation Definition ABSL animal biosafety level ACIP Advisory Committee on Immunization Practice Ag-ELISA antigen enzyme-linked immunosorbent assay (ELISA) AHF Argentine hemorrhagic fever AIDS acquired immunodeficiency syndrome ARDS adult respiratory distress syndrome AST aspartate aminotransferase ATA alimentary toxic aleukia AVA Anthrax Vaccine Adsorbed BSC biological safety cabinet BSL biosafety level BW biological weapons/biological warfare CBC complete blood count CCHF Congo-Crimean hemorrhagic fever CDC Centers for Disease Control and Prevention CF complement fixation CFR case fatality rate CFS chronic fatigue syndrome CIDRAP Center for Infectious Disease Research and Policy cm centimeter CNS central nervous system CSF cerebrospinal fluid CT computed tomography DFA direct fluorescence antibody 0701016F.doc x Abbreviations and Definition of Terms DIC disseminated intravascular coagulation DNA deoxyribonucleic acid DoD Department of Defense ECL electrochemiluminescence EDTA ethylenediaminetetraacetic acid EEE eastern equine encephalitis EIA enzyme immunoassay ELISA enzyme-linked immunosorbent assay EMG electromyography FAC free available chlorine FDA U.S. Food and Drug Administration g gram GI gastrointestinal HF hemorrhagic fever HFRS hemorrhagic fever with renal syndrome HI hemagglutination inhibition HIV human immunodeficiency virus IDSA Infectious Diseases Society of America IFA indirect fluorescent antibody IgG immunoglobulin G IgM immunoglobulin M IHC immunohistochemistry 0701016F.doc xi Abbreviations and Definition of Terms ABBREVIATIONS AND DEFINITION OF TERMS (CONT.) Abbreviation Definition IM intramuscular(ly) IND investigational new drug IV LAI LD50 LDH intravenous(ly) laboratory-acquired infection median lethal dose lactic dehydrogenase LVS live vaccine strain mL milliliter ng nanogram NP nasopharyngeal OP osopharyngeal PAPR powered air purifying respirator PBT pentavalent botulinum toxoid PCR polymerase chain reaction pfu/min plaque-forming units per minute PPE personal protective equipment PRNT plaque reduction neutralization titer RNA ribonucleic acid RT-PCR reverse transcription–polymerase chain reaction RUI report under investigation RVF Rift Valley fever SQ subcutaneous(ly) SEB staphylococcal enterotoxin B SIP Special Immunizations Program SOP standard operating procedure SST serum separator tube 0701016F.doc xii Abbreviations and Definition of Terms TBE tickborne encephalitis TERIS Teratogen Information System TT tiger top TU/kg Todd unit/kilogram USAMRIID U.S. Army Medical Research Institute of Infectious Diseases UV ultraviolet VEE Venezuelan equine encephalomyelitis VHF viral hemorrhagic fever VIG vaccine immune globulin VZV varicella zoster virus WBC white blood cell WEE western equine encephalitis WHO World Health Organization µg microgram µm micrometer 0701016F.doc xiii Chapter 1 - Introduction CHAPTER 1 INTRODUCTION PURPOSE OF THE MANUAL The purpose of this Occupational Health Manual for Laboratory Exposures to Select (BSL-3 & BSL-4) and Other Biological Agents is to serve as a reference for physicians in the Special Immunizations Program (SIP) at the U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID), Fort Detrick, Maryland. The manual is also intended to assist other health care providers who are called upon to evaluate USAMRIID employees who may have been exposed to bacterial agents, viral agents, and biological toxins being studied or tested in laboratories at USAMRIID. It is not intended as a definitive text on the medical management of illness resulting from exposure to specific agents. The manual represents the collective knowledge and experience of physicians in the SIP clinic. It also reflects the knowledge of experienced research scientists at the Institute, many of whom are national or international authorities on specific pathogens. BACKGROUND OF THE SIP The SIP dates back to 1942, when it was known as the Special Procedures Program. Its mission then (as now) was to vaccinate laboratory personnel at risk of exposure to biological and other agents. In the early 1970s, it became known as the Special Immunizations Program. The SIP clinic supports USAMRIID and extramural sites (i.e., outside USAMRIID) through administration of licensed and investigational vaccines, collection of safety and immunogenicity data on investigational vaccines, evaluation and reporting of adverse events, and oversight of the safety of SIP participants through annual evaluations, examinations, and laboratory testing. SUMMARY OF LABORATORY EXPOSURES AND ILLNESSES AT FORT DETRICK From 1943 to 1969, the United States conducted a biological weapons (BW) research and development program headquartered at Fort Detrick in Maryland. During this program, accidental infections or intoxications from potential warfare agents were periodically diagnosed among laboratory workers. The research in this program often involved production of high concentrations of organisms and experiments involving aerosolization of agents, which placed laboratory workers at a high risk for exposure and disease. A total of 423 laboratory-acquired infections (LAIs) were diagnosed until the program ended in 1969, with tularemia, brucellosis, Q fever, Venezuelan equine encephalitis (VEE), and anthrax, the five most commonly occurring infections [1,2]. Personal protective measures and improvements in safety practices were continuously implemented during the program as mechanisms of exposures were determined for laboratory incidents or accidents resulting in occupational illnesses. Even before the commercial availability of biological safety cabinets (BSCs) and development of vaccines, implementation of personal protective measures and safety practices prevented most infections from anthrax, glanders, and plague, most likely due to the higher infective dose required for infection with these agents [2]. However, infections continued to occur from agents having lower infective doses, such as Francisella tularensis, Brucella spp., and Coxiella burnetii. BSCs became commercially available in 1950 and were installed at USAMRIID from 1950 to 1954. Based on the risk of the research procedures, some laboratories were equipped with both Class I open- 0701016F.doc 1-1 Chapter 1 - Introduction fronted BSCs and Class III gastight BSCs. In one laboratory, the use of Class I and Class III BSCs by unvaccinated laboratory workers from 1954 to 1958 was associated with approximately two to four infections per year [1]. The exclusive use of Class III gastight cabinets by unvaccinated personnel dealing with similar organisms and conducting similar level-of-risk research during this same period was associated with only one infection per year [1]. Nearly all infections occurring in the two laboratories during this time were from organisms with low-infective doses, such as F. tularensis, Brucella, C. burnetii, and VEE virus. As investigational vaccines became available (i.e., live tularemia vaccine in 1959; VEE TC-83 vaccine in 1963; and Q fever vaccine in 1965), the number of infections further decreased, with only one or two infections occurring in each of the two laboratories in the subsequent 10 years. In a recent retrospective analysis, records in the SIP clinic of potential laboratory exposures to agents occurring at USAMRIID between 1989 and 2002 were reviewed for (1) agent exposure, (2) risk of exposure and disease (initial), (3) vaccination status, (4) management (post-exposure prophylaxis), and (5) outcome [3,4]. Exposures were classified as caused by bacteria, viruses, rickettsia, or toxins. Excluded from the review were 48 potential exposures to Herpes B virus and to 49 reports of either breaches in protection without potential exposure to a known agent or to Mycobacterium tuberculosis or individuals assessed for evaluation of community-acquired infections. A total of 250 individuals, 78% of whom were vaccinated prior to the potential exposure, were evaluated for potential exposures. The average number evaluated was 19 per year (approximately 400 at-risk individuals working in the laboratory during this time), and the range was 11 to 52 per year, with the majority being lower risk (assessed mainly as negligible or minimal risk of exposure). The large number of individuals evaluated can be attributed to USAMRIID’s policy of evaluating any potential risk of exposure or breach in protection occurring within biological safety level (BSL)-3 and BSL-4 laboratories. The number of persons evaluated per year varied widely, as a potential aerosolized exposure event required evaluation of all individuals within the area of aerosolization (as many as 16 persons for one exposure incident). However, the number of potential exposure events per se during this time remained relatively constant, with an average of 14.5 exposure events per year. During this period, only 5 individuals had infections diagnosed and 4 individuals had mild, self-limited symptoms after exposure to staphylococcal enterotoxins [4,5]. The confirmed infections included glanders, Q fever, localized vaccinia, chikungunya fever, and VEE [4]. Occupational illnesses in biodefense research are not expected to occur as frequently today as they did during the offensive BW program for several reasons. Biodefense research programs do not involve working with large volumes of organisms or frequent aerosolization experiments to the same extent as did the BW program. Also, laboratory practices and safety measures are more stringent, bringing improvements in engineering controls (i.e., directional airflow and better air-handling systems), advances in biosafety equipment (i.e., needleless systems), new vaccines, and applications of lessons learned from historical laboratory exposures. Increased worker awareness and modifications of laboratory practices and improvements in personal protective equipment may also contribute to the reduction in exposures and infections. Additionally, the current number of at-risk persons at USAMRIID (approximately 350 persons a year) is much lower than it was during the offensive BW program. However, an increase in the number of potentially higher-risk, animal aerosol-challenge efficacy experiments may be expected in future years, as the U.S. Food and Drug Administration’s (FDA’s) new “animal rule” for products whose efficacy cannot be tested in humans requires such testing. 0701016F.doc 1-2 Chapter 1 - Introduction In assessing the risk of disease in laboratory workers in today’s environment, the infective dose of the organism is an important factor, as demonstrated during the offensive BW program [2]. While most infections from agents with higher infective doses such as Bacillus anthracis, Yersinia pestis, and Burkholderia mallei were prevented even in a high-risk research setting with personal protective measures and safety education alone, two deaths from inhalation anthrax in unvaccinated individuals during the earlier BW program highlight the importance of vaccinating persons working with this agent. Improved control of infections from agents with lower infective doses (i.e., F. tularensis, C. burnetii, and VEE virus) in unvaccinated workers was observed when work with the agents was restricted to gastight Class III BSCs. However, unvaccinated laboratory workers employing BSL-3 or BSL-4 practices may still be at risk for percutaneous exposures or from unexpected aerosolization accidents (e.g., spills outside the BSC occurring when infectious agents are carried to and from Class III cabinets). Since as many as 80% of LAIs historically could not be attributed to a single exposure event (most LAIs were acquired by aerosol route and infection could not be associated with a single laboratory event) [6,7], vaccines may offer an added measure of protection for at-risk laboratory personnel. Vaccines, in some instances, may be preferable to the use of powered air purifying respirators (PAPRs), as PAPRs can restrict vision and manual dexterity, and thereby increase the risk of percutaneous exposures. While PAPRs may provide respiratory protection, percutaneous exposure may still occur (needlesticks alone are responsible for up to 25% of accidents in some laboratories). While protective measures such as Kevlar gloves and retractable needles may minimize percutaneous exposures, vaccines may afford added protection, particularly for agents for which post-exposure prophylaxis or treatment is not available (i.e., viral agents such as yellow fever). Data support vaccines as having contributed greatly to the decrease in infection rates. For example, a decrease in laboratory-acquired yellow fever was observed with the availability of the yellow fever vaccine [6,7]. Rates of typhoidal tularemia decreased from 5.7 cases to 0.27 cases per 1,000 at-risk employee-years with the introduction of a live, attenuated tularemia vaccine in the 1960s [8]. Guidelines for managing potential laboratory exposures at USAMRIID were developed, based on an analysis of recent laboratory exposures at USAMRIID from 1989 to 2002 and also from historical data at Fort Detrick [3]. The management guidelines were published in 2004 and included (1) information required in the history of the exposure event, (2) guidelines for assessing the risk of exposure by aerosol, percutaneous, mucocutaneous and cutaneous routes, (3) guidelines for assessing the risk of disease, and (4) recommendations for post-exposure prophylaxis in both vaccinated and unvaccinated individuals based on the risk assessment. PROTECTION FROM LABORATORY EXPOSURES AND PREVENTION OF ILLNESS AT USAMRIID USAMRIID’s success in limiting occupational illnesses in high-containment laboratories relies on the application of lessons learned from historical laboratory exposures as well as a comprehensive safety program and an SIP-managed vaccination and post-exposure prophylaxis program, which are described in the following sections. Comprehensive Safety Program The research laboratories at USAMRIID adhere to standards set by the Centers for Disease Control and Prevention (CDC), which include BSL-1 through BSL-4 [9]: 0701016F.doc 1-3 Chapter 1 - Introduction • • • • BSL-1, the basic level of containment, relies on standard microbiological practices. Work is restricted to defined and characterized strains of viable microorganisms not known to consistently cause disease in healthy adult humans. BSL-2 is appropriate for moderate-risk agents that cause human diseases by percutaneous or mucous membrane exposure or ingestion of infectious materials. BSL-3 is appropriate for indigenous or exotic agents with a potential for aerosol transmission and serious and possibly lethal infection. BSL-4 is restricted for work with dangerous and exotic agents that pose a high individual risk of life-threatening disease that may be transmitted by aerosol and for which no vaccine or therapy is available. Hazards to personnel are respiratory exposure to infectious aerosols, mucous membranes or broken skin exposure to infectious droplets, and autoinoculations. Access to BSL-3 and BSL-4 laboratories at USAMRIID is restricted to persons who have passed appropriate security background investigations and who carry appropriate electronic identification key cards and personal identification numbers. Furthermore, laboratory entrance and exits are tracked by security personnel. Laboratory workers receive focused medical evaluations as part of an annual medical surveillance program. They also receive extensive initial and annual education courses on general safety precautions, use of personal protective equipment (PPE), and emergency exit and decontamination procedures when working with the agents in their laboratories, including an extensive one-on-one mentoring program. Retraining is mandated based on procedures specific to any new studies. USAMRIID animal facilities also adhere to CDC BSLs for activities involving infectious disease work. The four levels of practices, safety equipment, and facilities are designated Animal Biosafety Levels 1, 2, 3, and 4 (ABSL-1 through ABSL-4) and provide increasing levels of protection to personnel and the environment that are comparable to those for BSL-1 through BSL-4 facilities [9]. Managed Vaccination and Post-Exposure Prophylaxis Program in SIP The laboratories at USAMRIID are housed as rooms within suites of laboratories [3]. The work in these suites is categorized according to type of agent: viral, bacterial, rickettsial, or toxin. This categorization allows for selection of specific vaccination recommendations for investigational new drug (IND) products and requirements for licensed products, e.g., anthrax vaccine (Figure 1-1). The vaccines are recommended or offered (IND products) on the basis of the specific agents stored or used within the suites to be accessed. To be vaccinated, individuals must be evaluated in the SIP, which involves providing a complete medical history and having a thorough physical examination, baseline laboratory tests, chest x-ray, and electrocardiogram. Criteria of immunogenicity of the vaccines and vaccine protocols are listed in Table 11. Admission to designated laboratories is based on pre-established criteria and based on potential risk for exposure. During their employment, individuals are followed regularly in the SIP clinic, where they receive annual physical examinations, are monitored for adverse events related to vaccination, and are given booster doses of vaccine as needed. 0701016F.doc 1-4 Chapter 1 - Introduction Figur e 1-1. FDA-Licensed and IND Pr oducts Available for At-r isk Labor ator y Wor ker s (Note: ACAAM smallpox vaccine now used in place of Dr yvax) Table 1-1. SIP Vaccines and Post-vaccination Immunogencity Criteria Vaccine Licensed by FDA Immunogenicty Criteria Anthrax Yellow fevera Yes Yes Japanese B encephalitis Yes Smallpoxa,b Yes Tularemiaa No Q Fever Venezuelan equine encephalomyelitisa,c (VEE) Western equine encephalitis (WEE) Eastern equine encephalitis (EEE) Rift Valley fever (RVF) Chikungunya fevera,f Tickborne encephalitisf Junin virusa,f No 3 weeks after third vaccine dose (given at 6 months) 1 month after vaccination One week after second vaccine dose (IXIARO) or after third vaccine dose (JE-VAX®) Evidence of a “take” (vesiculo-papular response); scab resolved (Days 21 to 28 after vaccination) Evidence of a “take” (vesiculo-papular response) by Day 28 and microagglutination titer ≥ 1:20 3 weeks after vaccination No PRNT80d titer ≥ 1:20e No No No No No No Pentavalent Botulinum Toxoid (PBT)g No PRNT80d titer ≥ 1:40e PRNT80d titer ≥ 1:40e PRNT80d titer ≥ 1:40e 4 weeks after vaccination 2 weeks after second vaccine dose 4 weeks after vaccination 4 weeks after the third dose of vaccine (or 4 weeks after a delayed booster) a b c d e f Live vaccine Booster doses given every 10 years, but every 3 years if working with monkeypox or Variola (work on Variola is conducted only at CDC). NOTE: The smallpox booster dose requirement for entry into or working with Variola in BSL-4 laboratories at the CDC was changed in 2009 by the Medical Advisory Board of the CDC Office of Health and Safety from an annual requirement to every 3 years booster dose requirement after review of new data. TC-83 vaccine is a live-attenuated VEE vaccine (primary vaccine); C-84 is an inactivated VEE vaccine (booster vaccine). 80% plaque reduction neutralization titer (PRNT) assay. Titer within 28 days after the primary series. No longer available at USAMRIID. 0701016F.doc 1-5 Chapter 1 - Introduction g PBT is losing potency, although serotypes A and B of Lot PBP-003 (lot currently used) still pass potency tests. The current CDC recommendation is to assume personal protective measures as the only protection against all five toxin serotypes and to have yearly booster doses (no titers measured) because of the decline in immunogenicity and potency. Antitoxin titers are no longer obtained in the protocol to verify immune response to the toxin serotypes. USER’S GUIDE TO MANUAL This manual is divided into five chapters. Chapters 1 and 2 include the Introduction and Risk Assessment of Occupational Exposures at USAMRIID, respectively. Chapters 3, 4, and 5 include information on the agents that are under study or that can affect researchers at USAMRIID. They are grouped under Bacterial and Rickettsial Agents, Viral Agents, and Biological Toxins. The information presented on each agent is as follows: Name of Agent • Nature of the Organism and Laboratory Exposures Nature of the Organism Experience with Laboratory Exposures BSL Working Conditions and Specific Laboratory Hazards • The Disease Clinical Features Diagnosis • Management of Potential Exposure to Agent (Asymptomatic Persons with Potential Exposure) Documenting the Exposure Diagnostic Testing of Asymptomatic Persons with Potential Exposure Post-exposure Prophylaxis • Management of Suspected Disease (Symptomatic Persons with Potential Exposure) Documenting the Exposure Diagnostic Testing for Symptomatic Persons with Potential Exposure Management of Clinical Specimens Treatment of Overt or Suspected Disease • Infection Control • References References 1. Wedum AG. The Detrick experience as a guide to the probable efficacy of P4 microbiological containment facilities for studies on microbial recombinant DNA molecules. J Am Biol Safety Assoc 1969;1:7-25. 2. Rusnak JM, Kortepeter MG, Hawley RJ, et al. Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers. Biosecur Bioterr 2004;4:281-93. 3. Rusnak JM, Kortepeter MG, Hawley RJ, et al. Management guidelines for laboratory exposures to agents of bioterrorism. J Occup Env Med 2004;46(8):791-800. 4. Rusnak JM, Kortepeter MG, Aldis J, Boudreau E. Experience in the medical management of potential laboratory exposures to agents of bioterrorism on the basis of risk assessment at the United States Army Medical Research Institute of Infectious Diseases (USAMRIID). J Occup Env Med 2004;46(8):801-11. 5. Rusnak JM, Kortepeter MG, Ulrich R, et al. Laboratory exposures to staphylococcal enterotoxin B. Emerg Infect Dis 2004;10:144-49. 0701016F.doc 1-6 Chapter 1 - Introduction 6. Pike RM. Laboratory-associated infections: incidence, fatalities, causes, and prevention. Ann Rev Microbiol 1979;33:41-66. 7. Sulkin SE, Pike RM. Viral infections contracted in the laboratory. N Engl J Med 1949;241(5):20513. 8. Burke DS. Immunization against tularemia: analysis of the effectiveness of live Francisella tularensis vaccine in prevention of laboratory-acquired tularemia. J Infect Dis 1977;135:55-60. 9. U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, and National Institutes of Health. Biosafety in Microbiological and Biomedical Laboratories, 5th ed. Washington DC, U.S. Government Printing Office, 2007. Only available at http://www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5toc.htm. Accessed December 12, 2007. 0701016F.doc 1-7 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID CHAPTER 2 RISK ASSESSMENT OF LABORATORY EXPOSURES AT USAMRIID DEVELOPMENT OF ASSESSMENT GUIDELINES Guidelines for assessment of laboratory exposure risk at USAMRIID were developed over several decades based, in part, on the clinical experience of laboratory workers who contracted disease after particular types of exposures. For example, in the late 1950s, 5 of 14 workers developed tularemia after an aerosol exposure resulting from dropping glass Petri dishes containing F. tularensis. As a result of this incident, studies of infectious bacterial aerosols were undertaken using nonpathogenic bacteria and cultures of air samples [1]. Results showed that bacterial aerosols resulting from dropping Petri dishes were produced from (1) rebounding of broken plates, (2) decontaminating the site with a spray apparatus, and (3) picking up broken Petri dishes. More aerosols were produced with the dropping of breakable glass Petri dishes than with unbreakable plastic Petri dishes. Dropping plastic Petri dishes resulted in reduced aerosolization to a range of just a few feet and a duration of a few minutes. Similarly, other studies using nonpathogenic bacteria and cultures of air samples demonstrated varying degrees of aerosolization with most laboratory procedures (i.e., pipetting, transferring of cultures using an inoculation loop, or withdrawing infectious agents from a sealed vial using a syringe and needle). Criteria to determine the exposure risk to infected animals were based on (1) clinical experience and (2) results of animal studies (i.e., studies on methods of cross-infection to other animals and excretion of organisms in the stool and urine) [2,3]. EVALUATION OF A POTENTIAL LABORATORY EXPOSURE OR ILLNESS AT USAMRIID According to USAMRIID policy, a physician must conduct an immediate evaluation of an individual who has: • • A potential laboratory exposure occurring within a containment suite or laboratory. A febrile illness with a temperature greater than 100.4°F and who worked in a containment suite within the past 1 to 3 weeks (depending on the incubation period of the biological agent). • An 8-fold rise in antibody titer to a biological agent. The stepwise process for evaluating a potential laboratory exposure is depicted in Figure 2-1. STEPWISE EVALUATION OF A POTENTIAL LABORATORY EXPOSURE The process for evaluating an asymptomatic employee who has a potential laboratory exposure is depicted on the left side of Figure 2-1. The process for managing a potential laboratory exposure can be separated into a series of steps [4], as follows: 1. Initial Evaluation. A physician conducts a detailed history of the exposure event, takes a medical history, and performs a medical exam. Baseline laboratory tests may be ordered, as indicated: − Baseline chemistries and complete blood counts (CBCs) are obtained on most individuals with minimal- or greater-risk exposures; baseline chest x-rays are taken if indicated. − Baseline serum for acute serologies may be drawn, frozen, and banked, if indicated. 0701016F.doc 2-1 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID Potential Laboratory Exposure OR Break in Laboratory Technique OR Febrile Illness OR 8-fold Rise in Antibody Titer* Step 1. Initial Evaluation • Evaluate if need for immediate isolation • Detailed history of exposure event or any break in laboratory technique • Baseline medical history and physical examination (H&P) • Baseline laboratory tests, as indicated Potential Laboratory Exposure (asymptomatic) Febrile Illness Step 2. Assessment • Assess risk of exposure (Tables 2-2 & 2-3) • Assess risk of disease (Table 2-4) • Assess need for postexposure prophylaxis (Figure 2-2) • Laboratory tests as indicated (i.e., baseline serology, nares culture/PCR) Possible Laboratory-Acquired Illness (LAI) Community-Acquired Illness Treat as indicated Step 3. Medical Management • Administer post-exposure prophylaxis or treatment and provide isolation/quarantine, as indicated and recommended for the specific agents • Follow-up in SIP clinic, (convalescent serologies and other laboratory tests, as indicated) • Formal written initial evaluation report Step 4. Corrective Actions Step 5. Maintenance of Records Documenting Potential Laboratory Exposure Event *If an 8-fold rise in antibody titer on evaluation by a physician may represent a potential LAI. Figur e 2-1. Stepwise Evaluation of Potential Labor ator y Exposur e 0701016F.doc 2-1 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID 2. Risk Assessment. The risk of exposure and the risk of disease are determined. Based on the risk assessment, the need for post-exposure prophylaxis, if available, is determined. Additional laboratory tests may be ordered, as indicated, to assess risk. − Reassess need for baseline laboratory tests and serum for acute serology. − Consider nares or throat cultures or PCR to further assess the risk of exposure to aerosolized agents. 3. Medical Management. If indicated based upon the risk assessment, and if available, postexposure prophylaxis should be given to exposed individuals. The requirement for isolation or quarantine should also be assessed. − The decision for isolation or quarantine, as well as the type of isolation required, will depend on the specific agent and the mode of transmission. Other elements that may be involved in determining the disposition of a patient may include: acuity of the illness, family assistance or other in-home supports, and the potential (or lack thereof) for adverse effects or risks to other persons in the home. − Monitoring and follow-up of most individuals generally will be performed in the SIP clinic. The frequency of follow-up clinical evaluations is based on the incubation period and virulence of the organism as well as the risk of disease from the exposure. Convalescentphase serologies are obtained at 2 weeks post exposure, if indicated. − A formal initial evaluation (see Appendix A) is written describing the incident in detail, the physical examination findings, the initial assessment of the risk of exposure and disease, the planned initial management of the individual based on the information known at the initial evaluation. 4. Corrective Actions. The Safety and Radiation Protection Office will review events related to the incident and propose corrective actions, as needed, to prevent a repeat occurrence. 5. Maintenance of Records Documenting Potential Laboratory Exposure Event. A formal final evaluation (see Appendix A) is written (generally 4 weeks or more after the original incident and includes (1) final assessment of the risk of exposure and disease based on the clinical course and laboratory test results and (2) corrective actions subsequently implemented. Reports of the initial and final evaluations of the potential exposure are maintained indefinitely. STEP 1: INITIAL EVALUATION During the initial evaluation, a detailed history of the potential laboratory exposure (Table 2-1), including the mechanism of exposure, agent(s) in use at the time of exposure, and type of protective equipment used, is documented. The information is obtained from the individual with the potential exposure, the individual’s supervisor and/or division chief, and biosafety personnel. Safety personnel, the individual’s supervisor and/or division chief, SIP physicians, and the Institute Commander must be notified within a few hours of the initial evaluation of the potential exposure. STEP 2: ASSESSMENT OF RISK Assessment of Risk of Exposure Based on the information obtained from the exposure incident, an assessment of risk of exposure to an infectious agent is based on the circumstances of the incident. The categorization of exposure risk is based on the likelihood of exposure occurring as a result of exposure to an agent [4] as follows: • • • Confirmed infection High (highly likely) Moderate (likely) 0701016F.doc 2-2 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID • • • Minimal (unlikely) Negligible (highly unlikely but unable to absolutely exclude) No risk (no greater risk than routine work in the laboratory with no breach in procedure occurring) Table 2-1. Documentation of a Detailed History of the Laboratory Exposure Event 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. Detailed description of exposure incident, how it occurred, who was involved. Duration of exposure risk, location (building, room), BSL of the laboratory. Types of procedures being conducted in the area at the time of the exposure, including whether aerosols were being generated (e.g., centrifugation). Infected animals in area with the potential to shed pathogens. Were animals ill? Pathogens in area at the time of the exposure (includes any agent used since the last decontamination of the laboratory). Route of exposure (i.e., inhalation, percutaneous, mucocutaneous, or ingestion). If percutaneous exposure, state whether exposure was from a needle or surgical blade, animal bite or scratch, or cut on sharp object. Was the needle sterile or did it have prior contact with a culture or solution containing the organism or an infected animal? Was the animal ill or not ill? If an animal was involved, consider exposure to other agents (e.g., Herpes B in rhesus macaque and mouth flora from animal bites). Time interval from incident to cleansing. What products were used for cleansing (e.g., Betadine, household bleach, or soap) and duration of cleansing? Were decontamination or other precautions taken in the area to prevent further exposure to other workers? Describe (i.e., decontamination of spill, closure of suite or area until appropriate decontamination could be performed, or repair of malfunctioning equipment). What protective equipment was in use by the individual at the time (type of gloves and number of pairs worn, type of mask or respirator, type of protective suit)? Was there a breach in the integrity of PPE (e.g., rip in protective suit or open zipper)? Were engineering controls (e.g., air-handling system, BSC, or decontamination showers) functioning properly? Source: Rusnak JM, Kortepeter MG, Hawley RJ, et al. [4] Table 2-2 includes examples of occurrences for high or moderate, minimal, negligible, and no risk for percutaneous, cutaneous, and mucosal exposures to infectious agents. Table 2-3 includes examples of risks for each of the four levels for aerosolized exposures to infectious agents. Because toxins are nonreplicating, exposures to them are categorized differently from exposures to infectious agents. For exposures to toxins, (1) high- or moderate-risk exposures are based on doses of toxin that are lethal or could cause severe symptoms; (2) minimal risk is based on doses that would cause mild, self-limiting symptoms; and (3) negligible risk is based on doses unlikely to cause symptoms. See individual sections on toxins. 0701016F.doc 2-3 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID Table 2-2. Risk Stratification of Percutaneous, Cutaneous, and Mucosal Exposures to Infectious Agents Risk Level High- or moderaterisk exposures Minimal-risk exposures Negligiblerisk exposures No-risk exposures Examples 1. Percutaneous exposure to any of the following: • Needle or surgical blade with prior contact with a solution or culture containing a viable agent or prior 2. 3. 1. 2. 3. 4. 5. contact with the blood, secretions, or tissue of an infected animal that is ill and likely to have organisms in the blood or secretions • Cuts near a centrifuge or from flasks containing a viable agent • Scratches or bites from an infected animal that is likely to be viremic or bacteremic (animal ill) Contact of a viable agent with nonintact skin (abrasions) or with the eye or mucous membranes Contact with a viable agent to intact skin but delay in cleaning the area Percutaneous exposure with a needle or surgical blade with prior contact to blood, secretions, or tissue of an animal infected with an agent, but animal not likely viremic/bacteremic or shedding organisms (animal not ill) Nonintact skin [abrasions] or eye or mucosal membranes contact with material unlikely to contain a viable infectious agent Direct contact of a viable infectious agent with intact skin and area cleaned immediately No discernible contact with a viable infectious agent or an infected animal (i.e., exposures to sterile needles or abrasion or cuts resulting from corner of desk or other objects highly unlikely to be contaminated in a BSL-3 or BSL-4 laboratory) Injury or breach in laboratory technique that did not occur in the presence of an animal or in a laboratory with an infectious agent or exposure to a substance later determined to be noninfectious Risk of exposure is no higher than usual when working in a laboratory (i.e., no breach in laboratory technique) 6. Source: Rusnak JM, Kortepeter MG, Aldis J, Boudreau E. [5] Table 2-3. Risk Stratification of Aerosolized Exposures to Infectious Agents Risk Level High- or moderaterisk exposures Exposure Criteria 7. While not wearing a respirator, direct splash of an infectious agent or aerosolization of a dried agent outside the BSC 8. While not wearing a respirator, exposure from centrifuge accident with viable organisms 9. Break in respiratory protection or inadequate air current in a BSL-4 suit (air entered the protective suit) in an environment with an infectious agent or infected animal, and where the agent is likely to be aerosolized 10. Exposure to culture not likely to aerosolize (i.e., dropping nonbreakable plastic culture plates and losing lids 11. Exposure from a splash or spill inside the BSC of an agent likely to be viable and without appropriate respiratory protection (respirator) 12. Exposure from a splash or spill outside the BSC of an agent unlikely to be viable and without appropriate respiratory protection (respirator) Minimal-risk exposures Negligiblerisk exposures No-risk exposures 13. Exposure to equipment outside the BSC that may have been exposed to agent when under the BSC (excluding a dried agent that could be easily aerosolized) 14. Break in respiratory protection in an environment with infectious agent or infected animals, but unlikely that the agent was aerosolized and the protective suit maintained an adequate air curtain (air did not enter into protective suit) 15. Break in respiratory protection in an environment with an infectious agent or infected animal, but the agent was contained in a BSC or the infected animal was unlikely to be shedding organisms, but this possibility cannot absolutely be excluded 16. Aerosolized exposure to a solution or liquid highly unlikely to be infectious Exposure to unknown material, later determined to be noninfectious (and no other identifiable risk factors for exposure) • A break in respiratory protection in an environment without an infectious agent or infected animals • No discernible increased risk than would normally occur from entering a biosafety laboratory (used appropriate personal protective measures and no breaches in laboratory technique) Source: Rusnak JM, Kortepeter MG, Hawley RJ, et al. [4] 0701016F.doc 2-4 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID Assessment of Risk of Disease from Infectious Agents The categorization of the risk of disease is based on the likelihood of disease occurring as a result of exposure to an agent [4] using the same categories to assess exposure risk (confirmed infection, high (highly likely), moderate (likely), minimal (unlikely), negligible (highly unlikely but unable to absolutely exclude), and no risk (no greater risk than routine work in the laboratory with no breach in procedure occurring). Assessment of disease risk (Table 2-4) includes characteristics of the disease and agent (lethal or infective doses of the agent and agent virulence), as well as host factors (whether the individual has been vaccinated against the agent, time since last vaccination, adequacy of last antibody titer results, or receipt of prophylactic antibiotics). Table 2-4. Assessment of Factors Influencing Risk of Disease after Exposure to an Agent What was the risk of disease? Note: Risk of disease is usually the same as risk of exposure or lower if individuals had prior vaccination, exposure to nonpathogenic strain, or was given antibiotic prophylaxis. Was there inhalational or mucosal contact or nonintact skin contact with agent? Was there immediate cleansing with disinfectant (time interval from incident to cleansing)? Immediate cleansing of agent may reduce disease risk. What was the estimated dose of exposure? What is the estimated infective dose/lethal dose of the agent? Was the individual vaccinated against the agent? Does the individual have protective antibody titers? How effective is the vaccine? Prior vaccination may lower the disease risk. What is the virulence of the organism? Exposure to nonvirulent strains may lower disease risk (i.e., nonvirulent Sterne strain of B. anthracis). Does individual have an illness or take medications that predispose for higher risk for disease? Are prophylactic antibiotics available against the organism? Post-exposure antibiotic prophylaxis may lower disease risk. Consider investigational antiviral agents to lower risk of disease in individuals with moderate- to high-risk exposures who are not vaccinated. 1. 2. 3. 4. 5. 6. 7. Source: Rusnak JM, Kortepeter MG, Aldis J, Boudreau E. [5] Assessment for Post-exposure Prophylaxis Management of exposed individuals with post-exposure prophylaxis will vary depending on the specific agent, the virulence of the agent, the route of exposure, the immune status of the individual, and the severity of illness, as well as on other criteria. As depicted in Figure 2-2, post-exposure prophylaxis is generally recommended for all moderate and high-risk exposures, regardless of immunization status, except for exposure to nonpathogenic organisms (i.e., Sterne strain of B. anthracis). • Most individuals with negligible risk exposures generally are not given post-exposure chemoprophylaxis, regardless of immunization status. • Vaccinated individuals with minimal-risk exposures have post-exposure prophylaxis decided on a case-by-case basis, but generally are not given antibiotics except for the following situations. Individuals with minimal-risk exposures that 1) involve potential percutaneous exposure (i.e., needlestick, bite, scratch) to an agent from handling a recently infected animal (infected usually within 2 to 4 weeks) that is not ill nor is likely to be bacteremic/viremic or 2) involve potential aerosol exposure to an infective agent from the dropping of unbreakable plastic culture plates outside a BSC (where aerosolization may occur but is minimal) are generally given post-exposure chemoprophylaxis regardless of immunization status. • Unimmunized individuals with minimal-risk exposures are generally given post-exposure chemoprophylaxis. • Most individuals with moderate or high-risk exposures are generally given post-exposure chemoprophylaxis, if available, regardless of immunization status. 0701016F.doc 2-5 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID Moderate/High Risk Exposures Others Percutaneous exposure to body fluids of recently infected (not ill) animal or Potential aerosol exposure from dropping culture plates and loss of lid Unvaccinated Antibiotics given to all exposures, regardless of vaccination status. a Negligible/No Risk Exposure Minimal Risk Exposure Antibiotics given to most exposures. a Vaccinated Antibiotic prophylaxis decided on case-by-case basis. a,b Antibiotics generally not given a Antibiotics are not given for exposures to nonpathogenic strains of B. anthracis in vaccinated individuals. Many percutaneous exposures in the laboratory from cuts on objects that are unlikely to be contaminated with infectious agents in vaccinated individuals are not given post-exposure antibiotic prophylaxis. Sources: Rusnak JM, Kortepeter MG, Hawley RJ, et al. and Rusnak JM, Kortepeter MG, Aldis J, Boudreau E. [4,5] b Figur e 2-2. Decision Tr ee for Recommending Post-exposur e Antibiotic Pr ophylaxis in Potential Exposur es to Bacter ial and Rickettsial Agents STEP 3: MEDICAL MANAGEMENT Most individuals evaluated for potential exposures can be managed on an outpatient basis. As most individuals with a potential exposure are evaluated within a few hours of the exposure and are asymptomatic, medical management generally consists of post-exposure prophylaxis (if available and if indicated) and clinical follow-up in the SIP clinic. An individual may be referred to a hospital, particularly if post-exposure therapy must be given intravenously or if the individual subsequently develops illness and requires treatment. Also, the need for isolation or quarantine must be reevaluated, particularly for individuals with minimal- or greater-risk exposures to agents that may result in severe infection and are contagious by the respiratory route or possibly to certain agents that are not endemic to the area. Particularly for higher-risk exposures to certain viral hemorrhagic fever agents, isolation or quarantine may be recommended in the BSL-4 containment suite at the Special Clinical Studies Unit (SCSU) at the National Institute of Health (NIH) in Bethesda Maryland (“Slammer” at USAMRIID is no longer operable as a BSL-4 containment suite for laboratory workers). Consultation with experts in infectious diseases and other specialties is recommended. Appropriate infection control measures must be implemented. Guidelines for diagnosis, post-exposure prophylaxis, treatment, and infection control measures are outlined in the chapters for the specific agents. A formal initial evaluation (Appendix A) is written describing the incident in detail, the physical examination findings, the initial assessment of the risk of exposure and disease, the planned initial management of the individual (i.e., decision concerning post-exposure prophylaxis and plans for follow-up) based on the information known at the initial evaluation. 0701016F.doc 2-6 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID STEP 4: CORRECTIVE ACTIONS All potential exposures will be reviewed by the Safety and Radiation Protection Office, and any corrective actions (i.e., new engineering or administrative controls, new recommendations for personal protective equipment, required safety training) that may help prevent recurrence of the exposure incident will be implemented. STEP 5: MAINTENANCE OF RECORDS DOCUMENTING POTENTIAL LABORATORY EXPOSURE EVENT In addition to the initial report documenting a detailed history of the exposure event, a final report on the outcome (generally completed at 4 weeks or longer following the original exposure event) is to be completed (Appendix A). The final report includes a final assessment of the risk of exposure and disease based on the clinical course and laboratory test results. The final report also includes any corrective actions that were implemented as a result of the incident. Both the initial and final reports are stored indefinitely. MANAGEMENT OF INDIVIDUALS WITH FEBRILE ILLNESS THAT MAY BE THE RESULT OF A LABORATORY EXPOSURE Individuals who present with a febrile illness must be evaluated to exclude a potential laboratory exposure. Guidelines for diagnosis, treatment, and infection control measures are outlined in the chapters for the specific agents. Isolation If the exposed individual works with an agent that may result in disease that may be spread by a respiratory route (i.e., SARS or avian influenza), the individual and clinic should ensure that appropriate infection control measures are followed to prevent secondary spread to coworkers and health care providers until an LAI can be excluded. Initial Evaluation, Assessment, and Laboratory Tests A medical history and physical exam should be performed. Also, a laboratory history should be obtained and include: • Agents in the laboratory with which the individual directly works as well as agents in the laboratory with which the individual does not directly work • Risk of potential exposure from the ongoing experiments • Any recent breaks in technique or protection. Information from the medical history and physical exam and absence of known breaks in protection while working in the laboratory will often provide insight into whether the illness is a community-acquired infection and not an LAI. Laboratory tests may be required to help confirm a community-acquired infection or confirm or exclude an LAI. Treatment Some individuals may need to be referred to a hospital for management, whereas other individuals may be managed on an outpatient basis. Appropriate consultation with experts in infectious diseases and other specialties is recommended. References 1. Barbeito MS, Alg RL, Wedum AG. Infectious bacterial aerosol from dropped Petri dish cultures. Am J Publ Health. 1961;27:318-22. 2. Wedum AG. Laboratory safety research with infectious aerosols. Publ Health Rep. 1964;79:619-33. 0701016F.doc 2-7 Chapter 2 – Risk Assessment of Laboratory Exposures at USAMRIID 3. Kruse RH, Wedum AG. Cross infection with eighteen pathogens among caged laboratory animals. Lab Anim Care. 1970;20:541-60. 4. Rusnak JM, Kortepeter MG, Hawley RJ, et al. Management guidelines for laboratory exposures to agents of bioterrorism. J Occup Env Med. 2004;46(8):791-800. 5. Rusnak JM, Kortepeter MG, Aldis J, Boudreau E. Experience in the medical management of potential laboratory exposures to agents of bioterrorism on the basis of risk assessment in the United States Army Medical Research Institute of Infectious Diseases (USAMRIID). J Occup Env Med. 2004;46(8):801-11. 0701016F.doc 2-8 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax CHAPTER 3 BACTERIAL AND RICKETTSIAL AGENTS ANTHRAX—Bacillus anthr acis Overview of the Laboratory Exposures Background Anthrax is an acute infectious disease caused by B. anthracis, a spore-forming, gram-positive bacillus, that is primarily a disease of herbivores, including cattle, sheep, goats, and horses. Humans generally contract anthrax when handling contaminated hair, wool, hides, flesh, blood, and excreta of infected animals and from manufactured products such as bone meal. Prior to development of a vaccine, those working in woolen mills were particularly at risk. Anthrax infection is introduced through scratches or abrasions of the skin, wounds, inhalation of spores, or eating insufficiently cooked infected meat. Based on nonhuman primate (NHP) studies, the inhalational lethal dose50 (LD50) is 2,500 to 55,000 spores [1]. Human exposures that occurred in the United States in 2001 to fine-grade aerosolized anthrax suggest that disease from B. anthracis can occur with inhalation of “one deep breath at site of release” [2]. However, most cases of anthrax have been cutaneous and occur on exposed parts of the body, e.g., hands, face, and neck. Experience with Laboratory Exposures Laboratory-acquired infections with B. anthracis have been primarily cutaneous anthrax resulting from exposures to organisms on the skin with or without a known history of an open skin lesion and have generally responded to medical treatment [3]. While cases of inhalational anthrax have occurred infrequently in the laboratory, the cases have been associated with a high rate of mortality even with medical treatment [4]. Laboratory-acquired gastrointestinal (GI) anthrax has not been reported. During the U.S. Biological Warfare (BW) program (1943–1969) at Fort Detrick (then called the U.S. Army Medical Unit, Fort Detrick), anthrax was one of the most commonly occurring laboratory infections. Between 1943 and 1945, 25 cases of cutaneous anthrax occurred, with the majority of lesions involving the forearms or hands (12 cases) or the lower facial area or neck (6 cases) [3,4]. The number of cases of anthrax markedly decreased in 1946 and 1947, coinciding with the pause in research that occurred after World War II. In 1946, a policy change at Fort Detrick required the use of long-sleeved operating gowns with gloves taped to the sleeves, replacing short-sleeved operating gowns previously used, to prevent cases of cutaneous anthrax. Upon resumption of anthrax research in 1948, anthrax cases occurred infrequently, with only two cases reported in the 5-year pre-vaccine period from 1948 to 1952 (including one fatal case of inhalational anthrax that occurred in 1951) [4]. This decrease in infections was attributed to changes in personal protective measures. After the introduction of the anthrax vaccine in 1952, only three cases of anthrax were diagnosed during the remaining 18 years of the BW program [4]. In one case, an individual who had received the initial three doses of the anthrax vaccine developed cutaneous anthrax just days before his scheduled 6-month vaccination (Dose 4). In a second case, an individual developed cutaneous anthrax a few days after receiving his third dose of anthrax vaccine at Month 6, instead of Week 4. In the third case, an unvaccinated electrician developed a lethal case of inhalational anthrax. The exact source of the inhalational exposure was undetermined despite an extensive epidemiological evaluation and led to the extension of anthrax vaccination of all craftsmen (plumbers, electricians, etc.) at Fort Detrick beginning 0701016F.doc 3-1 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax in 1958. No subsequent cases of anthrax have been diagnosed at the U.S. Army Research Institute of Infectious Diseases (USAMRIID) [5]. Specific Laboratory Hazards and Biosafety Level Working Conditions Anthrax spores may persist in the environment for prolonged periods of time (years) and are a source for infection in humans. Accidental parenteral inoculation or contact of the skin with B. anthracis from cultures or contaminated laboratory surfaces may result in infection. While cutaneous anthrax requires inoculation of the organism under the skin, most cases of cutaneous anthrax acquired in the laboratory had no known history of preexisting open skin lesions or trauma. Exposure to infectious aerosols (e.g., liquid spills of cultures) in the absence of proper respiratory protection may result in inhalational infection. B. anthracis may be present in blood, skin, lesion exudates, cerebrospinal fluid (CSF), pleural fluid, sputum, and, rarely, urine and feces of infected animals [6]. Biosafety Level-2 (BSL-2) facilities are recommended when working with B. anthracis cultures. BSL-3 facilities are recommended for work involving activities with a high potential for aerosol production. Animal Biosafety Level-2 (ABSL-2) facilities are recommended for studies using experimentally infected laboratory animals [6]. At USAMRIID, work with attenuated strains of B. anthracis is conducted in BSL-2 facilities. Animals experimentally infected with attenuated strains of B. anthracis are maintained in ABSL-2 facilities. All work with fully virulent B. anthracis is conducted in BSL-3 containment with enhancements (anthrax vaccination, mandatory clothing change, mandatory exit shower, HEPA filtration of exhaust air, and sewage sterilization). Animals experimentally infected with fully virulent strains of B. anthracis are maintained in ABSL-3 facilities with enhancements. All experimental aerosol studies of B. anthracis are conducted in a Class III BSC. An anthrax vaccine (Anthrax Vaccine Adsorbed [BioThrax® or AVA]), manufactured by Emergent BioDefense Operations Lansing Inc, Lansing, Michigan, is licensed by the FDA (Figure 3-1). It is derived from sterile culture fluid supernatant taken from an attenuated (nonencapsulated) strain. Until 2009, the vaccination series for preexposure prophylaxis consisted of six 0.5 mL subcutaneous (SC) doses at 0, 2, and 4 weeks; then at 6, 12, and 18 months, followed by yearly boosters [7]. Individuals were cleared for laboratory entry 3 weeks after receipt of the third dose of vaccine. In 2009, the FDA approved an amendment to modify both the route of administration and the dosing regimen of the anthrax vaccine primary series to be given as a series of five 0.5 mL intramuscular (IM) doses at 0 and 4 weeks, then 6, 12, and 18 months, followed by yearly boosters. This dosage regimen was associated with decreased local adverse events, attributed primarily to the change to the IM route of vaccine administration. Antibody concentrations with the new 5-dose primary series dosing regimen revealed lower titers at 8 weeks (4 weeks after Dose 2) compared to the previous dosing regimen (4 weeks after Dose 3) [8]. Individuals are still cleared for laboratory entry at 4 weeks after the receipt of Dose 2 (given at 4 weeks) of vaccine in the new 5-dose primary series, as it is standard procedure for workers to wear respiratory protection when laboratory work may result in generation of an aerosol. 0701016F.doc 3-2 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax Figur e 3-1. Licensed Anthr ax Vaccine for Pr e-exposur e Pr ophylaxis The Disease Clinical Features Human anthrax infection can occur initially in three forms—inhalational, cutaneous, or GI (including oropharyngeal disease)—depending on the route of exposure (Table 3-1). The incubation period following exposure to B. anthracis in unvaccinated individuals ranges from 1 to 7 days for GI anthrax and 1 to 12 days for cutaneous anthrax, but may be up to 43 days or longer for inhalational anthrax (inhaled spore forms may survive intact for 60 days or longer). As GI anthrax is unlikely to occur from a laboratory exposure due to current personal protective safety measures, further information in the manual will be provided mainly for cutaneous and inhalational anthrax. The major clinical feature of laboratory-acquired cutaneous anthrax is a painless local skin lesion [3]. The lesion may occur at the site of a preexisting skin lesion or abrasion or after a percutaneous injury. However, most cases of laboratory-acquired cutaneous anthrax had no history of an open skin lesion or trauma, but initially presented with a lesion described as a “pimple” that progressed into the typical cutaneous anthrax lesion. Anthrax lesions are often initially papular and then become vesicular before developing into an ulcer with the characteristic black eschar and localized edema (within 2 to 6 days). Lesions of laboratoryacquired cutaneous anthrax are most commonly located on the upper extremities and face. Individuals may also develop regional lymphadenitis and/or bacteremia if antibiotics are not initiated. 0701016F.doc 3-3 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax Table 3-1. Clinical Features of Anthrax Criterion Inhalational Cutaneous Route of Exposure Inhalation of aerosol Inoculation of skin Incubation Period 1 to 43 days or longer 1 to 7 days (up to 12) Prodrome of nonspecific febrile illness Often area pruritus, then, papule→ vesicle→ ulcer→ depressed black eschar Often brief improvement followed by abrupt onset of respiratory failure and hemodynamic collapse Clinical Presentation Chest x-ray findings: • Widened mediastinum • Pleural effusions • Pulmonary infiltrates may be observed • Associated hemorrhagic meningitis in 50% cases Mortality high if treatment initiated after onset of respiratory symptoms Prognosis Lesion painless and associated with localized edema Regional lymphadenitis common CFR <1% with treatment; 20% without treatment Gastrointestinal Ingestion of organism 1 to 7 days (usually 2 to 5) Abdominal pain, nausea, vomiting, and fever Bloody diarrhea, hematemesis, ulcers in GI tract, ascites Progression to acute abdomen or sepsis Oropharyngeal disease may present with fever, sore throat, difficulty swallowing, followed by ulcer/eschar and edema in oropharynx (may result in obstruction of upper airway) CFR 25% to 60% with/without treatment Meningitis Secondary seeding from bacteremia Variable Clinically indistinguishable from other etiologies of meningitis, except CSF is hemorrhagic in ≥50% of cases Gram stain may reveal grampositive bacilli Mortality high CFR was 86% in Sverdlovsk outbreak without treatment; CFR was 45% in 2001 in U.S. outbreak with treatment CFR = Case fatality rate During the BW program, lesions in 5 of the 25 cases of cutaneous anthrax diagnosed between 1943 and 1945 presented at sites of known preceding trauma [3,4]. The lesions in the other 20 cases presented initially as “pimples” without histories of known trauma. While cutaneous anthrax presented as a single lesion in most cases, 5 of the 25 cases presented with multiple lesions. Pruritus at the site was the earliest symptom, followed by a red macule that had developed into a red papular lesion and then into a vesicular lesion at the time of diagnosis. The lesion progressed from a red macule. The vesicular lesion either (1) ruptured and resulted in a small ulcer or (2) enlarged by direct extension or “a ring of satellite vesicles” (Figure 3-2) [3]. Localized edema, however, was only seen in 13 of the 25 cases on initial evaluation. Lesions were diagnosed early (within 48 hours of onset of the lesion in 13 of the 25 cases); therefore, all patients were initially afebrile on presentation. However, patients later commonly developed a fever (low-grade or high) of 2 to 6 days duration. Systemic symptoms of headache, malaise, or arthralgia were present in 20 of the 25 cases in varying degrees. A normal white blood cell (WBC) count at initial presentation was common, occurring in 22 of the 25 cases (subsequent leukocytosis was observed in only 10 cases). After initiation of antibiotics, the cutaneous lesions often progressed for 2 to 3 days, then 0701016F.doc 3-4 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax improved beginning with drying of the lesion and eschar formation, decreased edema, and decreased lymph node tenderness. The eschar often separated by Days 10 to 14 but was known to persist for longer periods (up to 67 days in 1 case). The defect remaining after eschar formation healed with secondary granulation. Figur e 3-2. Anthr ax Day 3 Skin Lesion Inhalational anthrax, on the other hand, generally presents as a prodrome manifesting as a nonspecific febrile illness (fever, malaise, fatigue, with/without nonproductive cough). After a brief period of improvement and generally within 2 to 4 days of the onset of symptoms, individuals may develop respiratory failure and sepsis. Chest x-ray findings of a widened mediastinum (may be subtle) and pleural effusions are common; pulmonary infiltrates may be observed. Approximately 50% of individuals with inhalational anthrax may develop hemorrhagic meningitis. Two cases of inhalational anthrax occurred during the BW program. The initial symptoms of the first case occurred in an unvaccinated bacteriologist who worked with B. anthracis and were nonspecific for inhalational anthrax. His symptoms consisted of a sore throat, malaise, rhinorrhea, and mild cough, which improved 2 days later but with subsequent tenderness over the maxillary sinus. The individual was treated with antibiotics for presumed sinusitis but presented on Day 4 of illness with a fever, respiratory distress, and slowed mental response. The second case was in an unvaccinated craftsman who presented in 1958 with a 1-day history of fever, severe headache, and myalgia with an injected throat and postnasal drip. The WBC count was normal, but the differential revealed a left shift. A subsequent chest x-ray showed a widened mediastinum, suggesting a diagnosis of inhalational anthrax that resulted in immediate admission and treatment. Whereas the first person was subsequently noted to have had a significant exposure to B. anthracis, the source of the exposure in the second person was unclear even after an extensive epidemiological investigation [4]. Differential diagnosis for cutaneous anthrax includes the following diseases/infections: • • • • Ulceroglandular tularemia (severe systemic toxicity usually uncommon) Staphylococcal or streptococcal cutaneous infections Orf (a viral disease acquired from sheep and goats; eschar formation and edema do not occur) Brown recluse spider bite (Loxosceles reclusa; may appear similar to anthrax lesion, but lesion is painful) Differential diagnosis for inhalational anthrax includes the following diseases/infections: 0701016F.doc 3-5 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax • • • • Pneumonia or influenza-like syndromes Pneumonic plague (hemoptysis common with plague but rare with inhalational anthrax) Tularemia (less likely to be fulminant) Q fever (usually asymptomatic or only mild illness) Diagnosis The most critical aspect in making a diagnosis of anthrax is a high index of suspicion associated with a compatible history of exposure. Cultures are considered the “gold standard” for anthrax diagnosis. In the initial 25 cases of cutaneous anthrax in the BW program, blood cultures were obtained only in 10 patients and were positive for B. anthracis in 3 of the patients. Gram stains performed on the lesions of the 25 cases were all positive for gram-positive bacilli, and cultures of the lesions were positive in 22 cases. While the Gram stains remained positive for as long as 5 days after initiation of antibiotics, cultures of the lesions were generally negative within 24 hours of initiation of antibiotics. Polymerase chain reaction (PCR)-based assays, fluorescent antibody assays (either indirect or direct fluorescent antibody assays [IFAs or DFAs]), or immunohistochemistry (IHC) may detect organisms in lesions where cultures are negative. Generally, serology is only useful in retrospective diagnosis [9]. Diagnostic sensitivity of the anti-PA (protective antigen) IgG assay is 97.8% and specificity is 97.6%. Serum should be collected at 0 to 7 days for acute-phase testing and at 14 to 28 days for convalescent-phase testing. Measurable antibodies in recent cases developed 1 to 16 days after onset of overt disease, but peak immunoglobulin G (IgG) levels may not be seen until 40 days after onset of symptoms. Antibody to protective antigen or the capsule develops in 68% to 93% of reported cases of cutaneous anthrax and in 67% to 94% of reported cases of oropharyngeal anthrax. However, serological testing may have limited use for anthrax diagnosis in laboratory workers who have received anthrax vaccine, as acute-phase serological testing will be positive from the anthrax vaccine. Other tests include IHC for detecting the organism in affected tissues and PCR-based assays for rapid identification of B. anthracis. PCR-based assays (as well as antigen-based assays) will initially become positive in the serum, often within 24 to 72 hours after aerosol exposure. Cutaneous Anthrax Cutaneous anthrax should be considered following development of a painless pruritic papule, vesicle, or ulcer—often with surrounding edema—that develops into a black eschar. Gram stain, culture, fluorescent antibody assays (IFA or DFA), or PCR tests of the lesion will usually confirm the diagnosis. Biopsy for histopathology and IHC may be considered. Blood cultures should be obtained as secondary bacteremia may occur, but generally blood cultures are negative early in the disease. Acute and convalescent serology should be obtained but will only confirm the diagnosis retrospectively. However, many laboratory workers studying B. anthracis have received the anthrax vaccine and will have positive acutephase serological tests due to vaccination, thereby further limiting the use of acute serological tests for diagnosis of anthrax in vaccinated laboratory workers. Inhalational Anthrax Diagnosis of inhalational anthrax is extraordinarily difficult, but the disease should be suspected with exposure to an aerosol containing B. anthracis. Early symptoms are nonspecific. However, development of respiratory distress in association with radiographic evidence of a widened mediastinum due to 0701016F.doc 3-6 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax hemorrhagic mediastinitis and presence of hemorrhagic pleural effusion or hemorrhagic meningitis should suggest the diagnosis of inhalational anthrax. A CT scan may be more sensitive than chest radiographs in detecting a widened mediastinum due to hemorrhagic mediastinitis [10,11]. Pulmonary infiltrates may also be observed in inhalational anthrax. Blood cultures are usually positive within 2 to 3 days of the onset of the illness, but serum PCR may be positive within 24 hours after an aerosol exposure. Diagnostic tests (i.e., PCR, culture, microscopy) should be performed on other specimens such as pleural effusions and CSF fluid, if present and/or clinically indicated. GI Anthrax Early diagnosis of GI anthrax may be difficult because of the rarity of the disease and its nonspecific symptoms. Only with a history of ingesting contaminated meat in the setting of an outbreak (or ingestion of spores) is the diagnosis usually considered. Blood cultures are usually positive within 2 to 3 days of illness. Stool may be cultured. Ascites, if present, may be obtained for Gram stain, culture, and PCR test. Meningitis Due to Anthrax Meningitis due to anthrax is clinically indistinguishable from meningitis due to other etiologies. An important distinguishing feature is that the CSF is hemorrhagic in up to 50% of inhalational anthrax cases. The diagnosis can be supported by identification of the organism (gram-positive rod) in CSF by microscopy and confirmed by culture or PCR. Management of Potential Exposures to B. anthracis (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to B. anthracis should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure (outlined in Chapter 2) are generally followed. Diagnostic Testing • Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater risk exposure generally require the following: – Two 10.0 mL serum separator tubes (SSTs) for research serology and storage at the time of exposure (should be within 7 days of exposure). (Serology may have limited value in individuals who have received the anthrax vaccine due to positive titers from the vaccine.) – Two 10.0 mL SSTs for convalescent research serology at 14 to 28 days post exposure, if indicated, for serologic diagnosis. – May consider (on case-by-case basis) nasal swab and/or throat swab for culture, immunofluorescent antibody, and PCR if within 24 hours of aerosol exposure. A positive nasal swab only indicates exposure to B. anthracis, and a negative nasal swab does not exclude exposure. Post-exposure Prophylaxis Males and Non-pregnant Females Post-exposure prophylaxis is indicated for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. Table 3-2 outlines chemoprophylaxis of persons with potential inhalational exposure to B. anthracis based on vaccination status. The optimal duration of post-exposure prophylaxis for cutaneous or GI anthrax without inhalational exposure is not known, but the 0701016F.doc 3-7 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax Advisory Committee on Immunization Practices (ACIP) suggests a duration of 10 to 14 days, because spore forms do not persist in these forms of the disease [7,9]. Table 3-2. Recommended Chemoprophylaxis for Potential Aerosol Exposure to B. anthracis Criterion Drug Regimen1 Immunized Ciprofloxacin, 500 mg PO, q12h, or Doxycycline, 100 mg PO, q12h2 60 days³ Duration Patient Followup Carefully monitor after discontinuation of antibiotics Nonimmunized, Vaccine Available Nonimmunized, Vaccine Unavailable or Not Given Ciprofloxacin, 500 mg PO, q12h, or Doxycycline, 100 mg PO, q12h Ciprofloxacin, 500 mg PO, q12h, or Doxycycline, 100 mg PO, q12h ≥60 days 60 days (and at least 30 days after Dose 3 received; vaccine dose for postexposure prophylaxis given as 0.5 mL SC at 0, 2, and 4 weeks after exposure)4 Carefully monitor after discontinuation of antibiotics Carefully monitor after discontinuation of antibiotics 1 Penicillins should not be used initially due to concern of resistance and due to low concentrations achieved with oral penicillin in pulmonary secretions, tissue, and within alveolar macrophages. (See section entitled “Pregnant females” for information on post-exposure prophylaxis during pregnancy.) Levofloxacin is FDA-approved for post-exposure prophylaxis for inhalational anthrax in adults 18 years of age and older. As safety data exist mainly for 28 days use, levofloxacin is recommended as a second-line drug for post-exposure prophylaxis. 2 Single dose of AVA IM may be recommended if ≥3 months since Doses 2, 3, or 4; or ≥6 months since Dose 5 or a booster dose. 3 The 30-day course of postexposure prophylaxis in fully vaccinated individuals is no longer recommended. The February 2009 draft of ACIP, the U.S. Department of Health and Human Services (HHS), and the Army Field Manual 8-284 (now in revision) recommend a 60-day course of post-exposure prophylaxis in persons who have been fully or partially vaccinated. 4 AVA administered post-exposure would be an investigational. The older AVA vaccine dosing regimen (3 SC injections given at 0, 2, and 4 weeks) are recommended for postexposure vaccination, due to earlier and higher levels of antibody titers with the older AVA dosing regimen [12]. Individuals with inhalational exposure to B. anthracis who are fully or partially vaccinated are recommended to receive chemoprophylaxis for 60 days (see Table 3-2). Individuals who completed the 5-dose primary series of the AVA and received their last vaccine dose more than 6 months ago and individuals who have received Doses 2, 3, or 4 of the vaccine more than 3 months ago should be given a booster dose to help ensure antibody titers do not decline to lower levels (see Table 3-2). Unvaccinated persons with potential aerosol exposure to B. anthracis may be given the anthrax vaccine as an IND because it is not a licensed therapy for post-exposure prophylaxis [12]. Post-exposure vaccination consists of three SC doses of vaccine given at 0, 2, and 4 weeks after exposure, due to the earlier and higher levels of antibody titers with the older AVA dosing regimen [12]. With post-exposure vaccination, post-exposure antimicrobial prophylaxis is recommended for at least 60 days [12]. Chemoprophylaxis for at least 60 days duration is recommended for individuals who are not receiving the anthrax vaccine. Pregnant Females Recommendation for antibiotic prophylaxis in pregnant women with significant risk exposure to B. anthracis: • Consultation with infectious disease and/or obstetrics experts or other experienced health care providers is highly recommended. Post-exposure prophylaxis must be assessed on an individual basis. Recommendation of post-exposure prophylaxis must weigh the risk of disease against the risk of receiving the post-exposure prophylaxis. 0701016F.doc 3-8 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax • • • If the strain of B. anthracis involved is known to be sensitive to penicillin, the drug of choice in pregnant women for post-exposure prophylaxis is amoxicillin. If the sensitivities of the organism are not known, prophylaxis with ciprofloxacin may be initiated and then switched to amoxicillin if the strain is found to be penicillin sensitive. Penicillin G procaine is FDA-approved for inhalational anthrax (post-exposure), but use of amoxicillin for this indication is “off-label” [12]. Ciprofloxacin is a Pregnancy Category C drug, and there are no adequate and well-controlled studies of its use in pregnant women. Although the Teratogen Information System (TERIS) concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Patients must be counseled concerning the risks and benefits of ciprofloxacin prophylaxis. Doxycycline is a Category D agent in pregnancy and may cause discoloration of fetus teeth if given during the last half of pregnancy [12]. The duration of post-exposure prophylaxis regimens during pregnancy is similar to recommendations in the absence of pregnancy (Table 3-2). Management of Suspected Anthrax Disease (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to B. anthracis must be recorded with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Blood for culture. • Whole blood in citrate tubes for PCR, antigen-based assays, and/or toxin assays (may use serum or citrated plasma as secondary source). • Serum for acute and convalescent serologies (limited value in immunized persons). • If skin lesion present, Gram stain, fluorescent antibody assays (IFA or DFA), PCR, culture of skin lesion, biopsy for histopathology, and IHC should be considered. • Chest x-ray (consider CT scan) to assess for widened mediastinum due to hemorrhagic mediastinitis of inhalation anthrax. • Pleural fluid, if present, for Gram stain, culture, and PCR (also cell count, lactic dehydrogenase [LDH], and protein). • Sputum, if present, for Gram stain, culture, PCR, and IFA (and specimens if diagnostic bronchoscopy performed). • CSF, if indicated, for Gram stain, culture, PCR, cell count, glucose, and protein. • Stool, if indicated, for culture. • Ascites, if present, for Gram stain, culture, PCR. Collection and Handling of Clinical Specimens Cutaneous Lesions Vesicular stage: Soak two dry sterile rayon or Dacron (not cotton) swabs in vesicular fluid from previously unopened vesicles. Note: Anthrax bacilli are most likely to be seen by Gram stain in the vesicular stage. Eschar stage: Perform Gram stain and culture of ulcer base or edge of eschar (rotate two moist (sterile, saline) rayon or Dacron (not cotton) swabs for 2 to 3 seconds at the base of the ulcer or underneath the 0701016F.doc 3-9 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax edge of the eschar. Transport in a sterile tube at room temperature (≤1 hour transport time) or at 2ºC to 8ºC (>1 hour transport time). Punch biopsy: Consider a punch biopsy for IHC testing if the patient has received antibiotics or has a gram-negative stain and culture despite a high index of suspicion for anthrax: • Tissue, fresh: 0.3 mm, store and transport frozen • Tissue preserved for histopathology: 0.3 mm, store and transport at room temperature • Biopsy of lesion preserved for histopathology: 0.3 mm, store and transport at room temperature (refrigerate if >1 hour transport time). Other Specimens Blood cultures: Collect appropriate blood volume and number of sets of blood cultures (usually two sets initially and then as clinically indicated). BACTEC™ cultures require 10 mL per bottle. If the patient has received antibiotics, blood cultures containing resin to absorb the antibiotics are recommended. Transport samples to the laboratory and hold at room temperature until they are placed on the blood culture instrument or incubator. Do not refrigerate. Blood for PCR: Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing and toxin assays. Transport chilled and not frozen. Collect one 7 mL serum tube or SST tube for serum for serologies (may refrigerate at 2°C to 4°C for 5 to 7 days; freeze at -30°C or lower if greater than 7 days). Sputum: If sputum is being produced, collect >1 mL of a lower respiratory specimen into a sterile container for Gram stain and culture or PCR. Inhalational anthrax usually does not result in sputum formation. Transport in sterile, screw-capped container at room temperature when transport time is <1 hour. For transport time >1 hour, transport at 2°C to 8°C. Nasal and throat swabs: Place material to be transported from 2 to 24 hours after collection in a transport container at 2°C to 8°C. Swabs (rayon or Dacron only, never cotton) should be moistened with the medium inside the packet, reinserted into the transport package, and transported at room or refrigerator (2°C to 8°C) temperature. Refrigerate at 2°C to 8°C if processing is delayed. Stool samples: Transfer ≥5 g of stool directly into a clean, dry, sterile, wide-mouth, leak-proof container. Transport unpreserved stool at room temperature when transport time is <1 hour. For transport time >1 hour, transport at 2°C to 8°C. CSF fluid: Obtain four tubes (in CSF kits) of CSF fluid with minimum of 2 to 3 mL per tube. Transport to laboratory at room temperature as soon as possible. Refrigerate if transport time >1 hour. Pleural fluid: Collect pleural fluid may be collected in a sterile syringe (or other sterile container if a larger volume). Transport to laboratory at room temperature as soon as possible. Refrigerate if transport time >1 hour. Treatment of Confirmed or Suspected Anthrax Inhalational Anthrax Clinical or subclinical meningitis in patients with inhalational anthrax is likely, based on the experience with 1979 Sverdlovsk outbreak (50% of 42 autopsies) and the 2001 outbreak (1 confirmed and 4 possible cases of meningeal involvement out of 11 cases). Therefore, treatment of inhalational anthrax and cases 0701016F.doc 3-10 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax with fulminant bacteremia should include a drug that penetrates the central nervous system. Treatment of inhalational anthrax should include ciprofloxacin (400 mg intravenous [IV] every 12 hours for adults) or doxycycline (200 mg IV loading dose, followed by 100 mg IV every 12 hours for adults) plus one or two additional drugs. Ciprofloxacin is recommended over doxycycline, as ciprofloxacin has higher penetration in the presence of meningeal inflammation than doxycycline [12,13]. The optimal combination of drugs for inhalational anthrax has yet to be determined, but infectious disease specialists currently suggest a combination of a quinolone, clindamycin, and rifampin for susceptible strains of B. anthracis [14]. Clindamycin is recommended in the therapy as the drug inhibits protein synthesis that may reduce the production of exotoxin (adult dose 900 mg IV every 8 hours). Riframpin has both CSF and intracellular penetration (adult dose 300 mg IV every 12 hours). Other drugs that may be considered include ampicillin or penicillin, meropenem, or vancomycin. Antibiotics should be adjusted pending results of sensitivity testing. Penicillin or other beta-lactam antibiotics should not be used as monotherapy for treating severe anthrax as resistance may develop due to inducible betalactamases. IV treatment should be switched to oral antimicrobial therapy when clinically appropriate. Early, aggressive drainage of pleural effusions (which may contain high levels of toxin and have mechanical effects that impair respiration) is recommended in inhalational anthrax patients and has been associated with decrease mortality. Also, individuals should complete a 60-day course of therapy as recommended for post-exposure prophylaxis for inhalational anthrax. [10, 12] (See Post-exposure Prophylaxis.) Consultation with an infectious disease physician is recommended. Anthrax immune globulin (AIG) derived from humans has been used as an investigational product for treatment of inhalational anthrax under an emergency IND protocol. Clinical data are insufficient at this time for the Conference Report on Public Health and Clinical Guideline for Anthrax to recommend use of AIG as a general recommendation for treatment of inhalational anthrax. The CDC continues to offer AIG under emergency IND on a case-by-case basis [12-15]. Cutaneous Anthrax Uncomplicated cutaneous anthrax (without inhalational exposure and without signs of systemic involvement) may be treated with shorter courses of oral antibiotics for 7 to 10 days duration [7,12]. Ciprofloxacin (500 mg tablet 2 times daily for adults), doxycycline (100 mg tablet 2 times daily for adults), or amoxicillin if penicillin sensitive (500 mg tablet 3 times daily for adults) are recommended therapies. If signs of systemic involvement or severe edema are present or if the lesion is located on the head or neck, IV ciprofloxacin and a multidrug approach as described in the inhalational anthrax treatment are recommended [12]. The edema associated with the skin lesion may be treated with nonsteroidal anti-inflammatory agents and is known to often progress for 1 to 2 days after initiation of treatment before improvement. If treated early, cutaneous anthrax in laboratory-acquired infections generally does not result in bacteremia [3]. In cases of cutaneous anthrax with systemic illness, the patient should be treated with IV antibiotics according to regimens used for inhalational anthrax. If aerosol exposure also occurred in individuals diagnosed with cutaneous anthrax, individuals should receive treatment for 60 days. Environmental Decontamination and Infection Control Anthrax spores germinate and form vegetative cells in environments rich in nutrients such as glucose, amino acids, and nucleosides. Vegetative bacteria generally survive poorly outside of mammalian hosts. 0701016F.doc 3-11 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax However, when in environments without nutrients, vegetative cells form spores that may survive in the environment for more than 40 years [16]. Studies on effective hand hygiene agents for removal of anthrax spores have used B. atrophaeus as a surrogate (based on reports that B. atrophaeus spores are slightly more resistant to germicides than are B. anthracis) [17]. These studies have found that hand washing with soap and water, 2% chlorhexidine gluconate, or chlorine-containing towels reduced the amount of B. atrophaeus spore contamination, whereas use of a waterless rub containing ethyl alcohol was not effective in removing the spores. Washes of 10, 30, and 60 seconds with either soap and water or chlorhexidine can eliminate 1.5 to 2.0 log10 CFU/mL of spores. Surface decontamination may be performed with a 10% solution of bleach (5% hypochlorite) for 30 minutes. Wiping with bleach is increasingly effective as wipe time increases. Health care workers should follow standard precautions when taking care of patients with infections associated with B. anthracis. As anthrax may be contracted from direct exposure to vesicle secretions of cutaneous anthrax lesions, gloves should be worn when examining possible anthrax lesions. Respiratory protection is recommended during autopsies. After an invasive procedure or autopsy, the instruments and area used should be thoroughly disinfected with a sporicidal agent (5% hypochlorite solution recommended). Person-to-person transmission of anthrax by a respiratory route does not occur. References 1. Inglesby TV, O’Toole T, Henderson DA, Bartlett JG, Ascher MS, Eitzen E, Friedlander AM, Gerberding J, Hauer J, Hughes J, McDade J, Osterholm MT, Parker G, Perl TM, Russell PK, Tonat K, Working Group on Civilian Biodefense: Anthrax as a biological weapon, 2002: updated recommendations for management, JAMA 2002; 287(17):2236–52. 2. Centers for Disease Control and Prevention: Update: investigation of bioterrorism-related inhalational anthrax—Connecticut, 2001, MMWR Morb Mortal Wkly Rpt 2001; 50(47):1049–51. 3. Ellingson HV, Kadull PF, Bookwalter HL, Howe C: Cutaneous anthrax: report of twenty-five cases, JAMA 1946; 131(14):1105–08. 4. Rusnak J, Kortepeter MG, Hawley RJ, Anderson AO, Boudreau E, Eitzen E: Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers, Biosecur Bioterr 2004; 2(4):281–3. 5. Rusnak JM, Kortepeter MG, Aldis J, Boudreau E: Experience in the medical management of potential laboratory exposures to agents of bioterrorism on the basis of risk assessment at the United States Army Medical Research Institute of Infectious Diseases (USAMRIID), J Occup Env Med 2004; 46(8):801–11. 6. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, and National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed¸ pp 122–5. Washington DC, U.S. Government Printing Office, 2007. 7. Advisory Committee on Immunizations Practices: Use of anthrax vaccine in the United States, MMWR Recomm Rep 2000; 49(RR-15):1–20. 8. Marano N, Plikaytis BD, Martin SW, Rose C, Semenova VA, Martin SK, Freeman AE, Li H, Mulligan MJ, Parker SD, Babcock J, Keitel W, El Sahly H, Poland GA, Jacobson RM, Keyserling HL, Soroka SD, Fox SP, Stamper JL, McNeil MM, Perkins BA, Messonnier N, Quinn CP, Anthrax Vaccine Research Program Working Group: Effects of a reduced dose schedule and intramuscular administration of anthrax vaccine adsorbed on immunogenicity and safety at 7 months; a randomized trial, JAMA 2008; 300(13):1532–43. 0701016F.doc 3-12 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax 9. Dewan PK, Fry AM, Laserson K, Tierney BC, Quinn CP, Hayslett JA, Broyles LN, Shane A, Winthrop KL, Walks I, Siegel L, Hales T, Semenova VA, Romero Steiner S, Elie C, Khabbaz R, Khan AS, Hajjeh RA, Schuchat A, Washington, D.C., Anthrax Response Team: Inhalational anthrax outbreak among postal workers: Washington, D.C., 2001, Emerg Infect Dis 2002; 8(10):1066–72. 10. Holty FE, Bravata DM, Liu H, Olshen RA, McDonald KM, Owens DK: Systematic review: a century of inhalational anthrax cases from 1900 to 2005, Ann Intern Med 2006; 144(4):270–80. 11. Jernigan JA, Stephens DS, Ashford DA, Omdenaca C, Topiel MS, Galbraith M, Tapper M, Fisk TL, Zaki S, Popovic T, Meyer RF, Quinn CP, Harper SA, Fridkin SK, Sejvar JJ, Shepard CW, McConnell M, Guarner J, Shieh WJ, Malecki JM, Gerberding JL, Hughes JM, Perkins BA, Anthrax Bioterrorism Investigation Team: Bioterrorism-related inhalational anthrax: the first 10 cases reported in the United States, Emerg Infect Dis 2001; 7(6):933–44. 12. Stern EJ, Uhde KB, Shadomy SV, Messonnier N: Conference report on public health and clinical guidelines for anthrax, Emerg Infect Dis 2008; 14(4):pii 07-0969. 13. Sejvar JJ, Tenover FC, Stephens DS: Management of anthrax meningitis, Lancet Infect Dis 2005; 5(5):287–95. 14. Gilbert DN, Moellering RC, Eliopoulos GM, Chambers HF, Saag MS, eds. The Sanford Guide to Antimicrobial Therapy 2010 (40th Edition). Antimicrobial Therapy, Inc, Sperryville, VA, 2010. 15. Walsh JJ, Pesik N, Quinn CP, Urdaneta V, Dykewicz CA, Boyer AE, Guarner J, Wilins P, Norville KJ, Barr JR, Zaki SR, Patel JB, Reagan SP, Pirkle JL, Treadwell TA, Messonnier NR, Rotz LD, Meyer RF, Stephens DS: A case of naturally acquired inhalation anthrax: clinical care and analyses of anti-protective antigen immunoglobulin G and lethal factor, Clin Infect Dis 2007; 44(7):968–71. 16. Manchee RJ, Broster MG, Stagg AJ, Hibbs SE, Patience B: Out of Gruinard Island, Salisbury Med Bull 1990; 68(Special Suppl):17–18. 17. Weber DJ, Sickbert-Bennett E, Gergen MF, Rutala WA: Efficacy of selected hand hygiene agents used to remove Bacillus atrophaeus (a surrogate of Bacillus anthracis) from contaminated hands, JAMA 2003; 289(10):1274–77. 0701016F.doc 3-13 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax Table 3-3. Diagnostic Testing for Anthrax Specimen Source and/or Test Aerobic blood cultures Antigen-based assays, PCR, and toxin assays Serum for acute and convalescent serologies Sputum for Gram stain, culture, PCR, and IFA Nasal/throat swab Eschar Gram stain, culture, and PCR Punch biopsy or scraping for IHC testing and histopathology 0701016F.doc Method of Collection Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Whole blood is preferable (collected in citrate tube); serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Collect 7–10 mL (minimum of 1 mL) of whole blood, serum, or citrate plasma for all assays Collect 7–10 mL (minimum 1 mL) in either SST or serum tubes and spun. Collect >1 mL of a lower respiratory specimen into a sterile container for Gram stain and culture. Use rayon or Dacron (not cotton) swabs. Moisten swab with medium inside the packet; reinsert into the transport packet. Rotate two moist (sterile, saline) rayon or Dacron (not cotton) swabs for 2 to 3 seconds at the base of the ulcer or underneath the edge of the eschar. Place in sterile tube. Obtain 0.3 mm fresh tissue specimen at advancing margin of lesion. Store in a sterile container. Transport Transport at room temperature. No refrigeration. Comments (if applicable) Further sets of blood cultures obtained as clinically indicated. Transport chilled (not frozen). Refrigerate at 2°C to 4°C for up to 7 days. Acute and convalescent serology at Day 0 to 7 and Day 14 to 28. Freeze at -30°C or lower if greater than 7 days. Retrospective diagnosis only. Acute serology may be of limited benefit in vaccinated persons who will have positive titers due to vaccination. Inhaled anthrax usually does not result in sputum formation. Transport in sterile, screw-capped container at room temperature, when transport time is <1 hour. Refrigerate (2°C to 8°C) for transport times >1 hour. Transport packet at room or refrigerator (2°C to 8°C) temperature. A positive swab only indicates exposure. A negative swab does not exclude exposure. Transport in a sterile tube at room temperature if ≤1 hour transport time. Anthrax bacilli less likely to be seen at eschar stage than vesicular lesion. Refrigerate (2°C to 8°C) if >1 hour transport time. Store and transport tissue specimen for histopathology at room temperature. Refrigerate if >1 hour transport time. Consider a punch biopsy for IHC testing and histopathology if the patient has received antibiotics and/or has a negative Gram stain and culture despite a high index of suspicion for anthrax. 3-14 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Anthrax Stool samples CSF for Gram stain, culture, cell count, glucose, protein, and PCR Pleural fluid for gram stain, culture, PCR, immunohistochemical stain, cell count, pH, LDH, protein, glucose 0701016F.doc Transfer ≥5 g stool directly into a clean, dry, sterile, wide-mouth, leak-proof container. Obtain 4 tubes of CSF fluid with 2–3 mL CSF per tube minimum for cultures and stains, PCR, chemistries, and cell count. Collect fluid in sterile syringe (or larger container if large amount of fluid). Transport unpreserved stool at room temperature when transport time is ≤1 hour and at 2°C to 8°C (refrigerated) if transport time is >1 hour. Transport tube to laboratory as soon as possible at room temperature. For GI anthrax only. Refrigerate if transport time >1 hour. Transport to laboratory as soon as possible (at room temperature if transport time is <2 hours). For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. 3-15 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague PLAGUE—Yer sinia pestis Overview of Laboratory Exposures Background Plague is an acute bacterial disease caused by the gram-negative bacillus Y. pestis (formerly known as Pasteurella pestis), resulting in bubonic (involving lymph nodes), septicemic, or pneumonic plague. Plague is normally transmitted from an infected rodent to a human by infected fleas or, uncommonly, as a result of direct handling of dead animal skins; by inhalation of aerosols from infected animal tissues; or by ingestion of infected animal tissues. The bite of an infected flea delivers between 25,000 and 100,000 Y. pestis organisms [1]. The infective aerosol dose is 100 to 500 organisms [2]. Experience with Laboratory Exposures Laboratory-acquired plague has been uncommonly reported in the medical literature, with only four cases reported from 1925 to 1962. During the U.S. BW program (1943–1969) at Fort Detrick (then called the U.S. Army Medical Unit, Fort Detrick), only one case of plague (non-lethal) occurred. In 1959, pneumonic plague was diagnosed in a laboratory worker, who likely acquired the infection during centrifugation of high concentrations of viable Y. pestis 5 days earlier [3]. One month prior to his infection, he had received his fourth dose of a killed, whole-cell plague vaccine that offered protection against bubonic plague but not pneumonic plague [4]. In 1993, a suspected case of cellulitis secondary to Y. pestis (an atypical presentation of plague) occurred at USAMRIID in a previously immunized individual who developed a painful 4 cm by 2.5 cm area of erythema and induration within 6 hours following a puncture on her hand from a needle contaminated with Y. pestis (syringe contained a high concentration of organisms). Symptoms resolved within 48 hours after initiation of doxycycline [5]. The low frequency of laboratory-acquired plague has been attributed to the higher infective dose of the organism and the instability of the organism in the environment. Laboratory-acquired plague cases from other laboratories has occurred most commonly from inhalational exposure [2,5]. Most infections occurred before the commercial availability of biosafety cabinets in 1950. While some cases were associated with poor safety practices that would not be tolerated in a laboratory today (e.g., rolling and then smoking a cigarette while working in the laboratory with Y. pestis), cases also occurred in laboratories with strict safety policies and in experienced researchers, such as the death of a scientist at Porton Down in 1962 who had worked with the organism for 12 years (specific source unknown) [3,6,7]. Seven deaths from laboratory-acquired plague were reported in the medical literature from 1898 to 1976 [6,8]. Two deaths in the United States from occupationally acquired plague have recently been reported. In 2007, a biologist for a National Park Service died from plague. It is believed that he acquired plague from aerosolization generated while performing a necropsy (in his garage) on a mountain lion. He did not wear protective gloves, mask, or other equipment because he believed the animal died from trauma (animal was subsequently confirmed to have plague). His occupational history was not documented in the medical records from his clinic visit one day following onset of symptoms (presumptive diagnosis of viral syndrome). The biologist was found dead at his home (2–3 days after onset of symptoms) [9]. In 2009, a scientist at the University of Chicago died from laboratory-acquired plague. The scientist had worked with an attenuated strain of Y. pestis (a vaccine strain) that was not known to cause disease in 0701016F.doc 3-16 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague healthy adults nor was required by the CDC to be studied under higher BSL conditions like virulent strains of Y. pestis. The scientist presented to the emergency department with a severe flu-like syndrome and breathing difficulties; he died within 12 hours [10]. The attenuated strain of Y. pestis was cultured from his blood. It is not known why this attenuated strain of Y. pestis caused disease in this individual (e.g., host factors, potential infectivity of agent even after prolonged artificial propagation) [10, 11]. Specific Laboratory Hazards and Biosafety Level Working Conditions Y. pestis may be present in bubo aspirates, blood, sputum, CSF, feces, and urine specimens from humans [12]. Laboratory personnel are at risk for aerosol exposure when infectious aerosols or droplets are generated during manipulation of cultures and during necropsy of infected animals [12]. Direct contact with cultures and infectious materials from humans or animals infected with Y. pestis or from bites of infected fleas also pose risks to laboratory workers. The organism is generally viable in low humidity, sunlight, and warm weather. BSL-2 facilities are recommended when working with potentially infectious Y. pestis cultures and materials, provided work does not result in aerosols. All work, including necropsies of infected animals, should be conducted in a BSC. BSL-3 containment and personal precautions are recommended for activities that result in droplet or aerosol production, for work with antibiotic-resistant strains, and for production of infectious materials [12]. At USAMRIID, work with attenuated strains of Y. pestis is conducted in BSL-2 facilities. Animals experimentally infected with attenuated strains of Y. pestis are maintained in ABSL-2 facilities. All work with fully virulent Y. pestis is conducted in BSL-3 containment with enhancements (mandatory clothing change, use of personal protective equipment, mandatory exit shower, HEPA filtration of exhaust air, and sewage sterilization). Animals experimentally infected with fully virulent strains of Y. pestis are maintained in ABSL-3 facilities with enhancements. All experimental aerosol studies of Y. pestis are conducted in a Class III BSC. The FDA-approved, killed, whole-cell plague vaccine that offered protection against bubonic plague is no longer manufactured and has not been available since 1999. The Disease Clinical Features The incubation period following Y. pestis exposure ranges from 1 to 8 days and is dependent on the route of exposure and the exposure dose. Plague can occur in three forms: bubonic, pneumonic, and septicemic (Table 3-4). Percutaneous exposures to Y. pestis may result in bubonic plague, which generally presents with a sudden onset of a febrile illness associated with tender lymph nodes. Inhalation of the organism may result in pneumonic plague, which presents as severe pneumonia that progresses rapidly to sepsis and death without appropriate antibiotic therapy. Septicemic plague results from either bubonic or pneumonic plague that has spread to the blood stream but may occur without preceding lymph node involvement. Septicemic plague presents with a febrile illness of sudden onset followed by septic shock and disseminated intravascular coagulation (DIC). 0701016F.doc 3-17 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague Table 3-4. Clinical Features of the Major Forms of Plague Criterion Route of Exposure Incubation Period Bubonic Percutaneous Inhalation of Y. pestis organisms or spread to lungs from septicemia 2 to 8 days 1 to 6 days Sudden onset headache, malaise, fever, and tender lymph nodes Sudden onset of headache, malaise, fever, myalgia, and cough Regional lymphadenitis (buboes) 25% of patients will have papule, vesicle, pustule, or furuncle at inoculation site Cellulitis reported but uncommon Clinical Presentation Pneumonic 80% can become bacteremic, causing hemorrhagic lesions in other lymph nodes and organs Liver and spleen often tender and palpable Septicemia, DIC, and shock possible Leukocytosis with a predominance of neutrophils CFR >60% without antibiotic therapy; <5% with therapy Prognosis CFR = Case fatality rate Cough with bloody sputum Pneumonia progressing rapidly to dyspnea and cyanosis Death from respiratory collapse/sepsis, usually within 2 to 5 days after illness onset Septicemic Bubonic or pneumonic plague that spreads to blood stream (may occur without preceding lymph node involvement) 1 to 4 days Sudden onset of fever, chills, headache, malaise, and myalgias followed by severe endotoxemia with shock and DIC May result in meningitis; spleen, liver, kidneys, skin, and brain most commonly affected organs Gram-negative rods (coccobacillus) on Gram stain or bipolar “safety pin” appearance of Wayson stain of sputum X-ray shows bronchopneumonia CFR nearly 100% if appropriate antibiotics not administered within 18 hours after onset of symptoms (approximately 50% case fatality rate in U.S. since 1950) CFR 30% to 50% with therapy; ~100% mortality if not treated within 18 hours after onset of symptoms Differential diagnosis of bubonic plague includes streptococcal or staphylococcal adenitis, tularemia, cat scratch disease, mycobacterial infection, primary genital herpes, or glanders. Differential diagnosis of pneumonic plague in the immunocompetent host would include inhalational anthrax, tularemia, community-acquired pneumonia, or Q fever (usually mild symptoms only). Differential diagnosis of septicemic plague includes all of the potential causes of septic shock. Diagnosis Initial diagnosis of plague is based primarily on clinical suspicion. A presumptive diagnosis can be made microscopically by identification of the coccobacillus in gram-negative, Wright-, Giemsa-, or Waysonstained smears from lymph node needle aspirate, sputum, blood, or CSF samples. The Wayson stain demonstrates a bipolar staining bacillus (light blue bacillus with dark blue polar bodies), giving the appearance of a “safety pin.” Immunofluorescent staining is also useful. Definitive diagnosis relies on culturing the organism from blood, sputum, CSF, or bubo aspirates. The organism grows slowly at normal incubation temperatures and may be misidentified by automated systems because of delayed biochemical reactions. It may be cultured on blood agar, MacConkey agar, or infusion 0701016F.doc 3-18 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague broth. Most naturally occurring strains of Y. pestis produce an F1-antigen in vivo, which can be detected in serum samples by immunoassay. A passive hemagglutination testing demonstrating fourfold rise in antibody titer in patient serum or a single titer ≥1:128 in an unvaccinated patient with a compatible illness is considered diagnostic (convalescent serum generally should be obtained 4 to 6 weeks later as early antibiotics may delay seroconversion for several weeks) [13]. PCR is not sufficiently developed for routine use, but it is a very sensitive and specific technique that is currently able to identify as few as 10 organisms per milliliter. A presumptive identification of Y. pestis can be made using PCR or antigen-capture enzymelinked immunosorbent assay [14,15]. Management of Potential Exposure to Y. pestis (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to Y. pestis should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure (outlined in Chapter 2) are generally followed. Diagnostic Testing • Individuals with negligible-risk exposures to Y. pestis generally only require observation. • Individuals with minimal- or greater risk exposure generally require the following: – Two 10.0 mL SSTs for research serology and storage at the time of exposure (should be within 7 days of exposure). – Two 10.0 mL SSTs for convalescent research serology at ≥14 days post exposure, if indicated, for serologic diagnosis. – May consider nasal swabs for culture (on a case-by-case basis) if within 24 hours of aerosol exposure. A positive nasal swab only indicates exposure to Y. pestis; a negative swab does not exclude exposure. Post-exposure Prophylaxis Male or Non-pregnant Females Post-exposure chemoprophylaxis should be initiated for asymptomatic persons exposed to Y. pestis and to health care workers and others with unprotected face-to-face contact with patients who have pneumonic plague. Chemoprophylaxis, which consists of doxycycline, 100 mg, orally every 12 hours; ciprofloxacin, 500 mg, orally every 12 hours; or tetracycline, 500 mg, orally 4 times daily, should continue for 7 days after last known or suspected Y. pestis exposure or until exposure has been excluded [16–18,19]. Chloramphenicol is also an alternative for post-exposure prophylaxis but has the risk of aplastic anemia. Trimethoprim-sulfamethoxazole may be considered for secondary-line therapy for post-exposure chemoprophylaxis but is known to be less effective than the primary prophylaxis regimens [19,20]. No chemoprophylaxis is required for face-to-face contact to individuals diagnosed with bubonic plague and without evidence of secondary pneumonic plague [20]. Pregnant Females Recommendations for antibiotic therapy in pregnant women with significant risk exposure to Y. pestis: • Consultation with infectious diseases and/or obstetrics (or other experienced health care provider) is highly recommended. Post-exposure prophylaxis must be assessed on an individual basis. 0701016F.doc 3-19 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague Recommendation of post-exposure prophylaxis must weigh the risk of disease against the risk of receiving the post-exposure prophylaxis. • Ciprofloxacin and trimethoprim-sulfamethoxazole are Pregnancy Category C drugs. Although TERIS (Teratogen Information System) concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Ciprofloxacin has been found to be effective in animal models in treating plague. Trimethoprimsulfamethoxazole has been considered a second-line drug for postexposure chemoprophylaxis. Doxycycline is a Category D agent in pregnancy and may cause discoloration of fetus teeth if given during the last half of pregnancy. Management of Suspected Plague (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to Y.pestis must be recorded with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Blood for cultures • Whole blood (citrate tubes) for F1-antigen assays (SSTs) and PCR (may use serum or citrated plasma as alternate secondary source) • Serum for immunoglobulin M (IgM) and IgG antibody titers for Y. pestis (acute and convalescent serology) • Sputum, lymph node aspirates, CSF (as indicated), and other fluids for Gram, Wayson, or fluorescent antibody stains and culture • All biopsies for histopathology and cultures/stains as stated earlier Collection and Handling of Clinical Specimens Sputum Transport lower respiratory tract (pneumonic) specimens for culture and Gram, Wright-Giemsa, Wright, or Wayson stain in sterile, screw-capped containers at room temperature for transport times less than 2 hours in length. For transport between 2 and 24 hours after collection, store specimens in a container and transport at 2ºC to 8ºC. Blood Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing; transport chilled and not frozen. Collect 7 mL of blood in SST or serum tube for serologies (may refrigerate at 2ºC to 4ºC for 5 to 7 days; freeze at -30ºC or lower if >7 days). Blood Cultures Collect appropriate blood volume and number of sets of blood cultures (usually 2 sets initially) and then as clinically indicated. BACTEC cultures require 10 mL per bottle (pediatric 5 mL per bottle). If the patient has received antibiotics, blood cultures containing resin to absorb the antibiotics are 0701016F.doc 3-20 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague recommended. Transport samples directly to the laboratory at room temperature. Hold them at room temperature until they are placed on the blood culture instrument or incubator. Do not refrigerate. Note: In suspected cases of plague, additional blood or broth culture (general nutrient broth) should be incubated at room temperature (22ºC to 28ºC), the temperature at which Y. pestis grows most rapidly. Do not shake or rock the additional broth culture so that the characteristic growth formation of Y. pestis can be seen clearly. Nasal Swabs Use rayon or Dacron® (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package and moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Aspirates or Biopsies Aspirates of involved tissue (bubo aspirates) or biopsied specimens (liver, spleen, lung) may yield little material; therefore, a sterile saline flush (1 mL sterile saline in a 10 mL syringe with 20 gauge needle) may be needed to obtain an adequate amount of specimen. Cap syringe and needle of aspirated sample, secure with tape, and send to the laboratory at room temperature (22ºC to 28ºC) for immediate processing. Refrigerate (2ºC to 8ºC) if processing will be delayed. Place tissues in sterile containers. Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. Treatment of Confirmed or Suspected Plague Pneumonic plague is invariably fatal if antibiotic therapy is delayed more than 1 day after onset of symptoms; therefore, early administration of antibiotics is critical. Antibiotics used in the treatment of plague include parenteral streptomycin, gentamicin, doxycycline, or chloramphenicol [16,19,21,22]. The patient is typically afebrile after the first 3 days of antibiotic therapy. Streptomycin has been considered the drug of choice (streptomycin 30 mg/kg/day administered IM in 2 divided doses). Due to ototoxicity or immediate nonavailability of streptomycin, gentamicin (5 mg/kg IV once daily or 2 mg/kg loading dose followed by 1.7 mg/kg IV every 8 hours) has been used as an alternate to streptomycin, either given alone or with doxycycline [21,22]. The dosage of aminoglycosides must be adjusted appropriately for renal insufficiency. In individuals with a contraindication to receiving aminoglycosides, doxycycline given as single-drug therapy may be administered either IV or orally (200 mg initially, followed by 100 mg every 12 hours). Tetracycline may be substituted for doxycycline, if doxycyline is not available. The recommended duration of treatment is at least 10 days (and at least 3 days following resolution of fever and other symptoms). Treatment may be changed from IV to oral therapy (generally doxycycline) when patient’s condition improves. Chloramphenicol, 25 mg/kg IV loading dose followed by 15 mg/kg IV 4 times daily for 10 to 14 days, is recommended for treating plague meningitis, pleuritis, or myocarditis (greater tissue penetration) [16]. Results obtained from animal studies indicate that quinolone antibiotics, such as ciprofloxacin and ofloxacin, may be effective. Recommended dosage of ciprofloxacin is 400 mg IV 2 times daily. Supportive therapy includes IV crystalloids and hemodynamic monitoring. Although low-grade DIC may occur, clinically significant hemorrhage is uncommon. Endotoxic shock is common, and supportive 0701016F.doc 3-21 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague care is required as needed with fluids and vasopressors. Finally, buboes rarely require any form of local care and generally recede with systemic antibiotic therapy. Incision and drainage pose a risk to others in contact with the patient due to aerosolization of the bubo contents. Aspiration is recommended for diagnostic purposes. Environmental Decontamination and Infection Control Y. pestis remains viable in water, soil, carcasses, hides, and grains for several weeks. At near-freezing temperatures, it will remain alive for months to years but is killed by 15 minutes of exposure to 55°C. It also remains viable for some time in dry sputum, flea feces, and buried bodies but is killed within several hours of exposure to sunlight and disinfectants. Aerosolized bacteria will survive up to 1 hour depending on atmospheric conditions. It also can persist up to 1 year in soil and 270 days in live tissue. Health care workers should use standard precautions for handling materials and patients with all forms of plague. Pneumonic plague may be spread person-to-person by droplet spread (droplet spread usually involves a distance of up to 6 feet from the person) [16,20]. Individuals with pneumonic plague should be isolated in a single room (does not require negative pressure). All persons entering the room should use respiratory protection. A surgical mask may be used for droplet precautions or a Technol N95 mask (or equivalent) [20]. Additionally, the use of gowns, gloves, and eye protection (face shield) for patient care should be implemented until the patient has completed 72 hours of antimicrobial therapy and is demonstrating clinical improvement and sputum culture is negative. References 1. Reed WP, Palmer DL, Williams RC Jr, Kisch AL: Bubonic plague in the southwestern United States: A review of recent experience, Medicine 1970; 49(6):465–86. 2. Franz DR, Jahrling PB, Friedlander AM, McClain DJ, Hoover DL, Bryne WR, Ravlin JA, Christopher GW, Eitzen EM: Clinical recognition and management of patients exposed to biological warfare agents. JAMA 1997; 278(5):399–411. 3. Burmeister RW, Tigertt WD, Overholt EL: Laboratory-acquired pneumonic plague. Report of a case and review of previous cases, Ann Intern Med 1962; 56:789–800. 4. Williams JE, Cavanaugh DC: Measuring the efficacy of vaccination in affording protection against plague, Bull World Hlth Organ 1979; 57(2):309–13. 5. Rusnak JM, Kortepeter MG, Aldis J, Boudreau E: Experience in the medical management of potential laboratory exposures to agents of bioterrorism on the basis of risk assessment at the United States Army Medical Research Institute of Infectious Diseases (USAMRIID), J Occup Environ Med 2004; 46(8):801–11. 6. Pike RM: Laboratory-associated infections: incidence, fatalities, causes, and prevention, Annu Rev Microbiol 1979; 33:41–66. 7. Death of Porton Scientist, Lancet 1962; 280(7253):463. 8. Pike RM: Laboratory-associated infections: summary and analysis of 3921 cases, Hlth Lab Sci 1976; 13(2):105–14. 9. Wong D, Wild MA, Walburger MA, Higgins CL, Callahan M, Czarnecki LA, Lawaczeck EW, Levy CE, Patterson JG, Sunenshine R, Adem P, Paddock CD, Zaki AR, Petersen JM, Schriefer ME, Eisen RJ, Gage KL, Griffith KS, Weber IB, Spraker TR, Mead PS: Primary pneumonic plague contracted from a mountain lion carcass, Clin Infect Dis 2009; 49:e33–8. 0701016F.doc 3-22 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague 10. Olivo A: CRPH: plague death not a threat to public health. Chicago Breaking News Center, September 20, 2009. Retrieved July 2, 2010 from http://www.chicagobreakingnews.com/2009/09/yersenia-pestis-malcolm-j-casadaban-university-ofchicago-medical-center-genetics-research-antibioti.html 11. Pike RM, Sulkin SE: Continued importance of laboratory-acquired infections, Amer J Pub Health. 1965; 55(2):190–9. 12. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed, Washington DC, U.S. Government Printing Office, 2007, pp. 158–60. Retrieved December 7, 2007 from www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5TOC.htm 13. Chu M: Laboratory manual of plague diagnostic tests, In: Centers of Disease Control and Prevention and World Health Organization, 2000. 14. Loïez C, Herwegh S, Wallet F, Armand S, Guinet F, Courcol RJ: Detection of Yersinia pestis in sputum by real-time PCR, J Clin Microbiol 2003; 41(10):4873–5. 15. Radnedge L, Gamez-Chin S, McCready PM, Worsham PL, Andersen GL: Identification of nucleotide sequences for the specific and rapid detection of Yersinia pestis, Appl Environ Microbiol 2001; 67(8):3759–62. 16. Inglesby TV, Dennis DT, Henderson DA, Bartlett JG, Ascher MS, Eitzen E, Fine AD, Friedlander AM, Hauer J, Koerner JF, Layton M, McDade J, Osterholm MT, O’Toole TO, Parker G, Perl TM, Russell PK, Schoch-Spana M, Tonat K for the Working Group on Civilian Biodefense: Plague as a biological weapon: medical and public health management, JAMA 2000; 283 (17):2281–90. 17. Russell P, Eley SM, Bell DL, Manchee RJ, Titball RW: Doxycycline or ciprofloxacin prophylaxis and therapy against experimental Y. pestis infection in mice, J Antimicrob Chemother 1996; 37:769– 74. 18. Centers for Disease Control and Prevention: Prevention of plague: recommendations of the Advisory Committee on Immunization Practice (ACIP), MMWR Recomm Rep 1996; 45(RR-14):1–15 19. Gilbert DN, Moellering RC, Eliopoulos GM, Chambers HF, Saag MS, eds. The Sanford Guide to Antimicrobial Therapy 2010 (40th edition). Antimicrobial Therapy, Inc, Sperryville, VA, 2010. 20. Kool JL. Risk of person-to-person transmission of pneumonic plague. Clin Inf Dis 2005;40:1166-72. 21. Boulanger LL, Ettestad P, Fogarty JD, Dennis DT, Romig D, Mertz G: Gentamicin and tetracyclines for the treatment of human plague: a review of 75 cases in New Mexico, 1985–1999. Clin Infect Dis 2004; 38(5):663–9. 22. Mwengee W, Butler T, Mgema S, Mhina G, Almasi Y, Bradley C, Formanik JB, Rochester CG: Treatment of plague with gentamicin or doxycycline in a randomized clinical trial in Tanzania, Clin Infect Dis 2006; 42(5):614–21. 0701016F.doc 3-23 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague Table 3-5. Diagnostic Testing for Plague Specimen Site and Test Method of Collection Transport Comments Plague Aerobic blood cultures Serum for F1 antigen assay and PCR acute Serum for convalescent serologies and serum IgM antibody 0701016F.doc Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Transport at room temperature. No refrigeration. Further sets of blood cultures obtained as clinically indicated. Also obtain general nutrient broth culture and incubate at room temperature (22°C to 28°C) at which Y. pestis grows more rapidly. Do not shake or rock the broth culture so that the characteristic growth formation of Y. pestis can be clearly visualized. Whole blood is preferable (collected in citrate tube) for PCR or antigen assays; serum (spun from serum collected in an SST or serum tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for all assays. Transport chilled (not frozen). Most naturally occurring strains produce an F1 Antigen in vivo, which can be detected in serum by immunoassay. Collect 1 mL minimum (7 mL adequate) of serum for serology in either SST or serum tubes and spun. Refrigerate at 2°C to 4°C for 5 to 7 days. PCR is presumptive identification only; not sufficiently developed for routine use but is very sensitive and specific (able to identify as few as 10 organisms/mL). Freeze at -30°C or lower if greater than 7 days. Acute and convalescent serology at Day 0 (within 7 days of exposure) and ≥ Day 14. (Passive hemagglutination testing demonstrating fourfold or greater rise in titer or ≥1:128 antibody titer to F1-antigen is considered diagnostic in unvaccinated patient with compatible illness. Convalescent titer should generally also be obtained 4 to 6 weeks later as early antibiotic treatment may delay serocoversion by several weeks.) 3-24 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Plague Collect >1 mL of a lower respiratory specimen into a sterile container for Gram stain and culture. Sputum for Gram stain, culture, fluorescent antibody stain, and PCR Transport in sterile, screwcapped container at room temperature, when transport time is <2 hours. For transport times between 2 to 24 hours, transport at 2°C to 8°C. Stains: Gram, Wright-Giemsa, Wright, or Wayson. Culture media: May grow on blood agar, MacConkey agar, or infusion broth. Caution: Organism grows slowly at room temperatures and may be misidentified by automated systems due to delayed biochemical reactions. PCR and antigen-capture ELISA are presumptive identification only. Nasal/throat swab Aspirates or biopsies CSF for Gram stain, culture, cell count, glucose, protein, and PCR 0701016F.doc Use rayon or Dacron (not cotton) swabs. Moisten swab with medium inside the packet; reinsert into the transport packet. Aspirates of involved tissue and biopsy specimens (i.e., aspirate bubo, liver, spleen, and lung) may yield little material; therefore, a sterile saline flush (1 mL saline in a 10 mL syringe with 20 gauge needle) may be needed to obtain an adequate amount of specimen. Transport packet at room or refrigerator (2°C to 8°C) temperature. See comments for sputum specimens for stains and culture. Aspirates: Syringe and needle with aspirated sample should be capped (recap using only one hand to prevent needle stick), secured by tape, and sent to lab at room temperature (22°C to 28°C) for immediate processing. Refrigerate (2°C to 8°C) if processing will be delayed. See comments for sputum specimens for stains and culture. Biopsies: Place tissues in sterile containers. Biopsies: Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. Obtain 4 tubes of CSF fluid with 2 to 3 mL CSF fluid minimum: Transport tube to laboratory as soon as possible at room temperature. • • • • tube 1 – cultures and stains tube 2 – PCR tube 3 – chemistries tube 4 – cell count See comments for sputum specimens for stain and culture. Refrigerate if >1 hour transport. 3-25 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia TULAREMIA—Francisella tularensis Overview of Laboratory Exposures Background Tularemia is caused by the gram-negative coccobacillus F. tularensis. Humans can acquire tularemia through contact of their skin or mucous membranes with the tissues or body secretions of infected animals (mostly rabbits and rodents such as hamsters and mice), from aerosolization of infective materials, and from bites of infected insects (deerflies, mosquitoes, or ticks). Less commonly, tularemia can be acquired through the ingestion of contaminated food or water. Infection can also be contracted from contaminated soil, grain, or hay, and from contaminated animal pelts and paws [1]. F. tularensis is highly infectious via the aerosol and intradermal routes, with as few as 10 organisms able to cause infection [2-4]. Person-to-person transmission of F. tularensis has not been documented. Experience with Laboratory Exposures Tularemia has been a commonly reported infection in research laboratories working with cultures of F. tularensis or with naturally or experimentally infected animals or their ectoparasites. Tularemia was the most frequent laboratory-acquired infection diagnosed during the period of the offensive BW program (1943 to 1969) at Fort Detrick (then called the U.S. Army Medical Unit, Fort Detrick) [3]. Van Metre and Kadull noted in 1959 that almost every individual who consistently worked with F. tularensis eventually acquired the disease [5]. Thirty-eight of 43 cases occurring between 1944 and 1956 involved individuals who worked directly with F. tularensis; however, the remaining 5 individuals (14%) had not worked with the organism and had a history of walking through the laboratory as the only risk factor [5]. Infections were mainly typhoidal or pulmonic and less commonly involved ulceroglandular or asymptomatic presentations. Even with the installation of BSCs in the early 1950s, infections in laboratory workers continued to occur, most likely related to the low-infective dose and the increased use of lyophilized preparations of the organisms, which are more likely to aerosolize [3]. The use of the phenol-killed Foshay tularemia vaccine during this period failed to prevent disease, but was associated with ameliorated symptoms from tularemia. From 1950 to 1959 (with use of the Foshay vaccine), Burke documented 73 cases of laboratory-acquired tularemia infections (6.92 cases/1,000 at-risk employee years). Typhoidal illness (both typhoidal and pulmonic tularemia) accounted for 60/68 (88%) symptomatic cases [6]. In 1959, the phenol-killed Foshay vaccine was replaced with a live, attenuated (live vaccine strain [LVS]) tularemia vaccine (Figure 3-3). During the following decade (1960 to 1969), only 11 patients were diagnosed with laboratory-acquired tularemia (incidence of 1.00 case/1,000 at-risk employee-years) [3,6]. Typhoidal disease decreased from 5.7 to 0.27 cases/1,000 employee years at risk. While there was not a statistically significant decrease in the incidence of ulceroglandular tularemia (0.76 to 0.54 cases/1,000 employee years at risk), there was a decrease in severity of disease and need for hospitalization for ulceroglandular tularemia cases in vaccinated individuals [6]. Nine of the 11 cases of tularemia diagnosed in the 10 year period after initiation of the live tularemia vaccine in 1959 occurred in vaccinated individuals [3,6]. Five cases of vaccine breakthrough involved ulceroglandular tularemia and occurred between 5 months to 10 years after vaccination, with tularemia agglutination titers ranging from 1:40 to 1:320 before the illness. While the live tularemia vaccine may 0701016F.doc 3-26 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia not have prevented ulceroglandular tularemia, the symptoms from ulceroglandular tularemia were ameliorated and individuals did not require hospitalization [3,6]. Four cases of asymptomatic or mild presumed typhoidal tularemia were diagnosed retrospectively in 4 vaccinated individuals (occurring from 2 to 6 years post vaccination) based on rises in serological titers and skin test seroconversion obtained routinely for surveillance purposes. Two of the four individuals recalled nonspecific illnesses in the recent interim period that may have represented mild typhoidal tularemia (recent past history of grippe with secondary bronchitis in one person and sore throat accompanied by chest and back pain in the other individual). Both individuals had been treated with tetracycline and symptoms resolved. The other two individuals denied having symptoms during the period of suspected infection. The two unvaccinated individuals who developed tularemia (1 case each of ulceroglandular and pulmonary tularemia) had a past history of pneumonic tularemia and persistently elevated antibody titers of 1:160 and 1:640 before exposure . The individual with recurrent pulmonary tularemia was engaged in pelleting, drying, milling, and packaging large quantities of the organism before his infection but denied any known break in physical protection. Figur e 3-3. IND Vaccine for Pr e-exposur e Pr ophylaxis for Tular emia Only three cases of tularemia were diagnosed in the biodefense research program from 1970 to 2010. Two cases were ulceroglandular and occurred in vaccinated individuals. In 1975, an individual presented with ulceroglandular tularemia that was believed to have been acquired through a break in the skin caused by a hangnail that was exposed after the removal of gloves while working with the organism. In 1980, an individual developed symptoms of ulceroglandular tularemia within 24 hours of a percutaneous exposure from a needle of a syringe that had contained 1 × 109 organisms. Both individuals were treated and recovered. In 2009, an unvaccinated individual presented with a febrile illness (headache, chills, body aches, dry cough, chest pain, nausea, mild diarrhea, and tenderness on medial side of the right eye orbit) and was ultimately diagnosed with pulmonic and probable oculoglandular tularemia. Her subsequent clinical course included development of a nodular yellow-pigmented lesion on the cornea of the medial right eye that was surrounded by injected conjunctiva, erythema nodosum (known to occur with tularemia), and vesiculopapular eruption with skin biopsy positive by culture and PCR for F. tularensis. Diagnosis was also confirmed by a ≥ 4-fold rise in tularemia microagglutination titers. The individual had not been vaccinated due to an elevated tularemia microagglutination titer of 1:40 and a history of a prolonged febrile illness several years earlier that could have represented symptoms from tularemia. The individual recovered after a short-term hospitalization and with antibiotic therapy (streptomycin and 0701016F.doc 3-27 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia ciprofloxacin). Investigation was unclear as to the exact source of the exposure, other than she had worked with F. tularensis in the laboratory. Specific Laboratory Hazards and Biosafety Level Working Conditions F. tularensis may be present in lesion exudates, respiratory secretions, CSF, blood, urine, and tissues from infected animals and fluids from infected arthropods [7]. Infection has been more commonly associated with cultures than with infected animals. In humans, between 10 and 50 organisms will cause disease if injected intradermally, whereas approximately 108 organisms are required with oral challenge. The human infectious dose by aerosol is approximately 10 to 50 organisms. F. tularensis Vaccine, Live, NDBR 101 is a live, attenuated vaccine available for workers at risk of exposure to F. tularensis in the SIP clinic under BB-IND 157. The vaccine has been associated with a decrease in cases of typhoidal and pulmonic forms of tularemia, and amelioration of symptoms in ulceroglandular laboratory-acquired tularemia [6]. Aerosol challenge tests in laboratory animals and human volunteers have also demonstrated significant protection compared to unvaccinated controls. While BSL-2 practices are recommended in clinical laboratories for activities involving clinical materials of human or animal origin suspected or known to contain F. tularensis (with notification of laboratory that the materials may contain F. tularensis), all manipulations of suspected cultures, preparatory work on cultures or contaminated materials for automated identification systems, animal necropsies, or experimental animal studies are recommended to be performed using BSL-3 or ABSL-3 practices, containment equipment, and facilities [7]. Characterized attenuated strains, such as F. tularensis Type B (LVS) and F. tularensis subsp novicida (strain U112) can be handled in BSL-2 facilities, but work with high concentrations of these strains should be performed under BSL-3 conditions [7]. At USAMRIID, work with attenuated strains of F. tularensis is conducted in BSL-2 facilities. Animals experimentally infected with attenuated strains of F. tularensis are maintained in ABSL-2 facilities. All work with fully virulent strains of F. tularensis is conducted under BSL-3 containment with enhancements (mandatory clothing change, use of personal protective equipment, mandatory exit shower, HEPA filtration of exhaust air, and sewage sterilization). Animals experimentally infected with fully virulent strains of F. tularensis are maintained in ABSL-3 facilities with enhancements. All experimental aerosol studies with F. tularensis are conducted in a Class III BSC. The live, attenuated LVS tularemia vaccine is administered intradermally with 15 pricks on the forearm. Individuals are then observed for a “take” as shown in Figure 3-4. A “take” is a reaction at the vaccine site, which is considered to be indicative of immunity from the vaccine. A typical “take” reaction is a papular lesion that progresses to a vesicle and then an eschar, although individuals may not experience all three stages. 0701016F.doc 3-28 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Figur e 3-4. Day 7 Reaction to Tular emia Vaccination The Disease Clinical Features Naturally-acquired tularemia presents primarily as ulceroglandular or typhoidal illness with or without pulmonary involvement, but also can present as glandular, oculoglandular, oropharyngeal, or a primary pneumonic form of disease, depending on the route of exposure. Laboratory exposure via aerosol to F. tularensis in general results in either pneumonic or typhoidal tularemia, whereas inoculation with organisms most likely results in ulceroglandular or glandular tularemia. Specific features of the six presentations are listed in Table 3-6. The average incubation period for all types of tularemia is 3 to 5 days (range is 1 to 14 days) [8] but has been reported to be as long as 21 days [9]. Tularemia typically lasts approximately 2 weeks. Most people respond rapidly to appropriate antibiotic therapy, with fever and generalized symptoms improving in 24 to 48 hours. Poor outcomes are associated with underlying comorbidity (alcoholism, diabetes), delay in seeking medical care, and delay in initiation of appropriate antibiotic therapy. Mortality of untreated pneumonic tularemia can approach 60%. During the BW program, the diagnosis of typhoidal tularemia included symptoms of typhoidal tularemia with or without pulmonic involvement. Pulmonic involvement was noted in 11 of the 34 individuals with typhoidal tularemia from 1944 to 1956 [5]. Five of these 11 individuals had no evidence of pneumonia on physical exam. An analysis of 42 cases of typhoidal tularemia over the following 4 years (1956 to 1959) showed similar incidence, severity, and type of respiratory symptoms in individuals regardless of the presence or absence of pulmonic involvement [10]. Seventeen of the 42 individuals (40%) had abnormal chest x-rays, with 15 individuals having a single, oval pneumonic lesion with indistinct borders and a density ranging from 2 to 8 cm. Three individuals had multiple densities, and one individual had diffuse bronchopneumonia and lobar pneumonia. Hilar adenopathy was observed in 9 cases and pleural effusions in 5 cases. An unpublished animal study suggested that the presence of single versus multiple nodules on a chest x-ray may be related to the infective dose, with single pulmonic lesions occurring after small challenge doses and multiple infiltrates occurring after larger challenge doses [11]. 0701016F.doc 3-29 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Most cases of typhoidal tularemia include fever and malaise as the most prominent symptoms [5], often associated with localized symptoms suggesting a respiratory tract infection, including a nonproductive to mildly productive cough, pharyngeal irritation or injection, and rhinitis. Many individuals described their throat pain as having a “raw feeling in the throat.” Chest pain was also observed in some but not all individuals and was described as a vague substernal chest tightness or pleuritic in nature [10]. Moderate hepatomegaly was observed in 3 cases (7%) and moderate splenomegaly in 5 cases (9%) in Van Metre’s analysis of patients. WBC counts ranged from 5,000 to 10,000 cells/mm3 at the time of diagnosis in 75% of cases, with 20% of cases presenting with leukocytosis and WBC counts between 10,100 to 12,000 cells/mm3 [5]. Secondary cutaneous lesions may occur in 8% to 35% of cases, with the most commonly reported skin lesions being papular or macular-papular lesions, vesicular or pustular lesions, erythema nodosum, and erythema multiforme [12,13]. Six of the 9 cases of laboratory-acquired ulceroglandular tularemia in one cohort resulted from percutaneous exposures (needle, broken glass, autopsy tool) and only 3 cases were without a history of trauma [5]. All lesions occurred on the hand or digits, beginning as a papulo-erythematous lesion with progression to ulceration in 3 cases. Systemic symptoms of fever and malaise were absent, moderate, or severe, but all individuals exhibited localized tender regional lymphadenopathy. Initial WBC count at diagnosis ranged from 4,100 to 9,400 cells/mm3 [5]. The lesions of all individuals responded rapidly to antibiotics with defervescence within 2 days and rapid regression. Differential diagnoses of the various presentations of tularemia are as follows: • • • • • • Ulceroglandular tularemia—anthrax, Pasteurella infections, primary syphilis, rat-bite fever, rickettsial pox, and scrub typhus Glandular tularemia—bubonic plague, cat-scratch disease, mycobacterial infection, and streptococcal or staphylococcal adenitis Typhoidal tularemia—brucellosis, disseminated mycobacterial or fungal infection, Q fever, viral infections that cause a mononucleosis syndrome (i.e., Epstein-Barr virus, cytomegalovirus, and acute human immunodeficiency virus [HIV] infection), or rickettsial infections, prior to the onset of rashes Oculoglandular tularemia—adenoviral infection, cat-scratch disease, herpes simplex, syphilis, or tuberculosis Oropharyngeal tularemia—streptococcal pharyngitis, infectious mononucleosis, and adenoviral infection Pneumonic tularemia—community-acquired pneumonia, inhalation anthrax, pneumonic plague, Q fever, or viral pneumonia 0701016F.doc 3-30 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Table 3-6. Clinical Features of the Major Forms of Tularemia Criterion Route of Exposure Incubation Period Types of Tularemia Ulceroglandular Glandular Typhoidal Acquired when organisms enter by intradermal route (often through unapparent breaks in skin). Acquired mainly from an intradermal route (often through unapparent breaks in the skin), but no ulcer formed. Occurs mainly after inhalation of infectious aerosols but can occur after any mode of transmission (intradermal or GI). Acquired when Acquired from ingestion organism enters of the organism. conjunctiva, either from rubbing eyes with contaminated fingers or from splashes or aerosol. Can be primary after inhalation of organisms or secondary after hematogenous spread from other sites. Average 3 to 5 days (range 1 to 21 days) Average 3 to 5 days (range 1 to 21 days) Average 3 to 5 days (range 1 to 21 days) Average 3 to 5 days (range 1 to 21 days) Average 3 to 5 days (range 1 to 21 days) Average 3 to 5 days (range 1 to 21 days) 75% to 85% of naturally occurring cases. 5% to 10% of naturally occurring cases. < 5% to 15% of naturally occurring cases. 1% to 2% of naturally occurring cases. < 5% of naturally occurring cases. Presents with photophobia, increased lacrimation, painful conjunctivitis, and edema of the eyelid. Symptoms mainly confined to throat (fever, severe throat pain). Pulmonary involvement in > 80% of typhoidal cases and 30% of ulceroglandular cases or can present initially as primary pneumonia. Presents with ulcerated skin lesion in 3 to 5 days with painful lymphadenopathy Presents with fever and fever, chills, headache, cough, and and tender lymphadenopathy myalgia. but no skin lesion. Lesion is a red and painful papule that necrotizes, resulting in a painful ulcer 0.4 to 3 cm in diameter with irregular raised border. Signs and Ulcer may appear before, during, or Symptoms after lymphadenopathy. Dark scab or eschar may resemble that of anthrax and cover area of ulceration. Affected lymph nodes 0.5 to 10 cm in diameter, usually tender and may become fluctuant, rupture, and create draining sinus tracts in skin. Presents with fever, prostration, headache, malaise, sore throat, and weight loss. No adenopathy, but may have anorexia, nausea, vomiting, and diarrhea. Pneumonia from secondary pleuropulmonary involvement. Nonproductive cough or substernal chest discomfort. Oculoglandular Small yellowish conjunctival ulcer. Localized tender adenopathy of preauricular, submandibular, or cervical nodes. Oropharyngeal Acute exudative or membranous pharyngotonsillitus where necrosis and suppuration may occur. Pneumonic Presents as an atypical pneumonia with fever, headache, sore throat, cough with minimal or no sputum production, substernal tightness, and with or without pleuritic chest pain. Can have focal necrosis and lesions as with ulceroglandular form. Organisms may disseminate via hematogenous spread to multiple organs, and sepsis can occur. 0701016F.doc 3-31 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Prognosis CFR as high as 60% if untreated. CFR may vary with subsepecies of F. tularensis (Type A more severe disease versus Type B rarely fatal), clinical form of tularemia (higher mortality in pneumonic and typhoidal forms of tularemia), delay of treatment, or comorbidities (i.e., alcoholism, diabetes). CFR = Case fatality rate 0701016F.doc 3-32 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Diagnosis Tularemia can be diagnosed by recovery of F. tularensis in culture or from serologic evidence of infection in a patient with a compatible clinical syndrome. Although the organism is difficult to culture, it can be recovered from blood, ulcers, lymph nodes, conjunctival exudates, sputum, pleural fluid, gastric washings, and pharyngeal exudates. The organism will NOT grow on most routine solid media or gramnegative selective media such as MacConkey agar, as the organism requires cysteine or cystine for growth. Cultures should be subcultured onto glucose cysteine agar plates. The diagnosis of typhoidal tularemia was made by culture in 28 of 38 laboratory workers between 1956 and 1959 with gastric, pharyngeal, and sputum cultures being the main sites for diagnosis [10]. Due to the infective nature of the organism, the clinical laboratory should be alerted when cultures from patients with suspected tularemia are being requested. Most diagnoses of tularemia are made serologically using a bacterial microagglutination assay or enzymelinked immunosorbent assay (ELISA). However, vaccination status and previous infection can affect assay results. Measurable levels of antibodies that agglutinate F. tularensis appear within 1 week of infection, but levels high enough to allow confidence in specificity of the serologic diagnosis (an agglutination titer > 1:160, for example) do not appear until more than 2 weeks after infection [14]. Older studies at Fort Detrick indicated that titers did not peak until around 7 weeks after exposure. More recent studies show that titers are usually negative at Week 1, positive by the end of Week 2, and peak after Week 4 or 5. The serologic response may be blunted by prior administration of antibiotics. Because F. tularensis may cross-react with Brucella, Proteus OX19, and Yersinia organisms and because detectable antibody levels may persist for many years after a bout of tularemia, the serologic diagnosis of an acute infection should, ideally, be made only if a fourfold or greater increase in serological response is seen during the course of a patient’s illness. DFA stain confirmation may be performed in tissue, sputum, or culture. PCR of clinical specimens (blood, ulcer exudates, etc.) has been investigated as an alternative and is a promising diagnostic modality. Further testing with clinical specimens is required before the exact role of PCR in the diagnosis of tularemia cases can be established. Management of Potential Exposure to F. tularensis (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to F. tularensis should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure (outlined in Chapter 2) are generally followed. Diagnostic Testing Note: Due to the infective nature of the organism, the clinical laboratory should be alerted when cultures from patients with suspected tularemia are being requested. • Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater-risk exposure generally require the following: – Two 10.0 mL SSTs for research serology and storage at the time of exposure (should be within 7 days of exposure). – Two 10.0 mL SSTs for convalescent research serology at ≥ 14 days post-exposure, if indicated, for serologic diagnosis. Antibody titers to tularemia generally do not appear until ≥ 2 weeks after exposure and peak at week 4 or 5. 0701016F.doc 3-33 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia – May consider (on a case-by-case basis) nasal swabs for epidemiological purposes (if within 24 hours of aerosol exposure) for culture, DFA, and PCR (depending on route of exposure, if known). A positive nasal swab only indicates exposure to F. tularensis, and a negative swab does not exclude exposure. Post-exposure Prophylaxis Post-exposure prophylaxis is indicated for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. Because the incubation period for tularemia is usually 3 to 6 days and immunity following vaccination takes about 2 weeks to develop, postexposure vaccination with the investigational LVS is not considered a viable strategy to prevent disease. Males and Non-pregnant Females • Post-exposure chemoprophylaxis should ideally be initiated within 24 hours of exposure and continued for at least 14 days. Tetracycline prophylaxis for shorter durations was ineffective in preventing tularemia in humans [15]. Recommended chemoprophylaxis for tularemia is doxycycline, 100 mg, orally 2 times daily or ciprofloxacin, 500 mg, orally, 2 times daily for 14 days. • A nonhuman primate study demonstrated that streptomycin (bactericidal against tularemia) for 5 days was effective in preventing tularemia [13]. However, aminoglycosides are not currently recommended for postexposure prophylaxis for tularemia. Pregnant Females Recommendations for antibiotic prophylaxis in pregnant women with significant risk exposure to F. tularensis: • Consultation with infectious diseases and/or obstetrics (or other experienced health care provider) is highly recommended. Post-exposure prophylaxis must be assessed on an individual basis. Recommendation of post-exposure prophylaxis must weigh the risk of disease against the risk of receiving the post-exposure prophylaxis. • Ciprofloxacin is a Pregnancy Category C drug. Although the TERIS concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Doxycycline is a Category D agent in pregnancy that may cause discoloration of fetus teeth if given during the last half of pregnancy. Management of Suspected Tularemia Disease (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to F. tularensis must be recorded with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing Note: Due to the infective nature of the organism, the clinical laboratory should be alerted when cultures from patients with suspected tularemia are being requested. • • • Blood for culture (with subculture onto glucose cysteine agar plates). Whole blood in citrate tubes for PCR (may use serum or citrated plasma as a secondary source). Serum for IgM and IgG agglutination (microagglutination assays) or ELISA titers (acute and convalescent serologies). 0701016F.doc 3-34 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia • • • If skin lesion or lymph node aspirate, Gram stain and culture of skin lesion and biopsy for histopathology, PCR, and DFA. Sputum or pleural fluid, if present, for Gram stain, culture (onto glucose cysteine agar plates), DFA, and PCR. Other tissues or exudates, as indicated, for culture, stain, DFA, PCR, and histopathology. Collection and Handling of Clinical Specimens Blood Cultures Collect appropriate blood volume and number of sets of blood cultures (usually 2 sets initially and then as clinically indicated). BACTEC adult cultures require 10 mL per bottle. If the patient has received antibiotics, blood cultures containing resin to absorb the antibiotics are recommended. Blood cultures should be maintained in a closed system, and clinical isolates from blood or any other site should be handled under BSL-3 conditions. Transport directly to laboratory at room temperature. Hold at room temperature until placed on the blood culture instrument or incubator. Do not refrigerate. Cultures should be subcultured onto glucose cysteine agar plates. Skin Lesions Biopsied tissue or scraping of an ulcer is preferable, but a swab of an ulcer is an acceptable alternative. Submit tissue or scraping in a sterile container. Transport at room temperature for immediate processing at USAMRIID. If processing is delayed, specimen should be kept refrigerated (2ºC to 8ºC). Biopsies or Lymph Node Aspirates Collect biopsy, scraping, swab, or aspirate of ulcer, lymph node, or other involved tissue. For biopsy, place the tissue or scraping in sterile container. Transport at room temperature for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Serum Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes for PCR (may use serum or citrated plasma as alternate secondary source); transport chilled and not frozen. Collect one 7.0 mL red top or TT tube for serum for serology during week 1 of illness following exposure (may refrigerate at 2ºC to 4ºC for 5 to 7 days; freeze serological specimens at -30ºC or lower if > 7 days). Collect convalescent serum ≥ 14 days after exposure (antibodies appear in most patients by 2 weeks after onset and peak at 4 to 5 weeks). Nasal Swabs Use rayon or Dacron (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package and moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Other Specimens Other respiratory specimens (pleural fluid, sputum), gastric washings, or corneal scrapings can be sent for culture. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. 0701016F.doc 3-35 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Treatment of Confirmed or Suspected Tularemia Antibiotic treatment for tularemia infection is critical. Individuals who are not treated may have an illness that lasts for months and is characterized by malaise, weakness, and weight loss. Treatment may include the following regimens: • Streptomycin (7.5 to 10 mg/kg every 12 hours for 10 to 14 days, not to exceed 1 g IM twice daily as doses greater than 2 g streptomycin per day do not increase efficacy) or gentamicin 5 mg/kg/day in divided doses (gentamicin may also be given once daily at a dosage of 5 mg/kg/day, but not as well studied) [16–19]. The addition of chloramphenicol to streptomycin also may be considered in cases of meningitis [18]. Note: Gentamicin is not an FDA-approved treatment for tularemia. Dosage of aminoglycosides needs to be adjusted for renal insufficiency. Streptomycin is associated with ototoxicity that may be irreversible. • Alternatives to treatment are 1) doxycycline (100 mg IV twice daily), which has a higher rate of relapse than treatment with aminoglycosides (only bacteriostatic), for 14 to 21 days duration (may switch to oral therapy when patient clinically improved) or 2) ciprofloxacin (400 mg IV twice daily) for 14 to 21 days (may switch to oral ciprofloxacin 750 mg every 12 hours when patient is clinically improved) [18]. Chloramphenicol (15 mg/kg IV four times daily; maximum dose of 4 grams/day) for 14 to 21 days has been used for treatment of tularemia but generally is no longer used due to the risk of aplastic anemia. • • Lymph nodes should be needle-aspirated or surgically drained if fluctuant. Environmental Decontamination and Infection Control The organism can remain viable for weeks in water, soil, carcasses, hides, and grains and for years in frozen rabbit meat. It is resistant for months to freezing temperatures, but it is easily killed by heat (55°C for 10 minutes) and disinfectants. Person-to-person transmission of tularemia from a respiratory route does not occur; therefore, neither isolation nor quarantine is necessary. Standard precautions are appropriate for care of patients with draining lesions or pneumonia (including disinfection of soiled clothing or bedding, contaminated equipment, etc.). References 1. Heymann DL, ed: Control of Communicable Diseases Manual, 18th ed, pp 573–76. Washington DC, American Public Health Association, 2004. 2. Franz DR, Jahrling PB, Friedlander AM, McClain DJ, Hoover DL, Bryne WR, Pavlin JA, Christopher GW, Eitzen EM Jr: Clinical recognition and management of patients exposed to biological warfare agent, JAMA 1997; 278:399–411. 3. Rusnak J, Kortepeter MG, Hawley RJ, Anderson AO, Bourdrea E, Eitzen E: Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers, Biosecur Bioterr 2004; 2(4):281–93. 4. Pike RM: Laboratory-associated infections: summary and analysis of 3,921 cases, Health Lab Sci 1976; 13(2):106–14. 5. Van Metre TE Jr, Kadull PJ: Laboratory-acquired tularemia in vaccinated individuals: a report of 62 cases, Ann Intern Med 1959; 50(3):621–32. 6. Burke DS: Immunization against tularemia: analysis of the effectiveness of live Francisella tularensis vaccine in prevention of laboratory-acquired tularemia, J Infect Dis 1977; 135(1):55–60. 0701016F.doc 3-36 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia 7. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, and National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed, pp 137–39. Washington DC, U.S. Government Printing Office, 2007. www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5TOC.htm. Accessed December 7, 2007. 8. Centers for Disease Control and Prevention: Emergency preparedness and response—Tularemia. http://www.bt.cdc.gov/agent/tularemia. Accessed February 11, 2008. 9. Darling RG, Woods JB, ed: USAMRIID’s Medical Management of Biological Casualties Handbook, 5th ed. USAMRIID, Fort Detrick, Maryland, 2004. 10. Overholt EL, Tigertt WD, Kadull PJ, Ward MK, Charkes ND, Rene RM, Salzman TE, Stephens M: An analysis of forty-two cases of laboratory-acquired tularemia: treatment with broad spectrum antibiotics, Am J Med 1961; 30:785–806. 11. Overholt EL, Tigertt WD: Roentenographic manifestations of pulmonary tularemia, Radiology 1960; 74:758–65. 12. Syrjälä H, Karvonen J, Salminen A: Skin manifestations of tularemia: a study of 88 cases in northern Finland during 16 years (1967–1983), Acta Derm Venereol 1984; 64(6):513–16. 13. Eliasson H, Bäck E: Tularemia in an emergent area in Sweden: an analysis of 234 cases in five years, Scandinavian J Infect Dis 2007; 39:880–89. 14. Evans ME, Friedlander AM: Tularemia, Chapter 24 in Sidell FR, Takafugi ET, Franz DR, eds: Textbook of Military Medicine: Medical Aspects of Chemical and Biological Warfare, Part I, pp 503–12. Washington DC, Office of the Surgeon General at TMM Publications, Borden Institute, 1997. 15. McCrumb FR Jr, Snyder MD, Woodward TE: Studies on human infection of Pasturella tularensis; comparison of streptomycin and chloramphenicol in the prophylaxis of clinical disease, Trans Assoc Am Phys 1957; 70:74–79. 16. The choice of antibacterial drugs, Med Lett Drug Ther 1998; 40(1026):33–42. 17. Sanders CV, Hanh R. Analysis of 106 cases of tularemia. J La State Med Soc. 1968;120:391-3. 18. Gilbert DN, Moellering RC, Eliopoulos GM, Chambers HF, Saag MS, eds. The Sanford Guide to Antimicrobial Therapy 2009 (39th Edition). Antimicrobial Therapy, Inc, Sperryville, Virginia, 2009. 19. Dennis DT, Inglesby TV, Henderson dA, Bartlett JG, Ascher MS, Eitzen E, et al. Consensus statement. Tularemia as a biological weapon. Medical and public health management. JAMA 2001; 285:2763-73. 0701016F.doc 3-37 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Table 3-7. Diagnostic Testing for Tularemia Specimen Site and Test Method of Collection Transport Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Transport at room temperature. Whole blood is preferable (collected in citrate tube) for PCR; serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Store at 4ºC until tularemia can be ruled out (not frozen). PCR is investigational. Refrigerate at 2ºC to 4ºC for 5 to 7 days. Acute and convalescent serology at Day 0 and Day 14 or greater; antibodies appear in most patients by 2 weeks after onset and peak at 4 to 5 weeks after onset. Do not refrigerate. Aerobic blood cultures Serum for PCR Comments 1 Further sets of blood cultures obtained as clinically indicated. Culture media: The organism will NOT grow on most routine solid media or gram-negative selective media (i.e., MacConkey agar) as the organism requires cysteine or cystine for growth. Cultures should be subcultured onto glucose cysteine agar plates (may grow in thioglycolate broth, chocolate agar, modified ThayerMartin medium, buffered charcoal-yeast agar, or cysteine heart agar with 9% chocolatized sheep blood). Some isolates may grow on blood agar initially but not on subpassage. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for PCR. Collect 1 mL minimum (7 mL adequate) of serum for serology in either red top or TT tubes and spun. Serum for acute and convalescent serologies Freeze at -30ºC or lower if greater than 7 days. Caution: Diagnosis of acute infection should also include 4fold rise in serology as detectable antibody levels may persist for many years after tularemia and antibodies may also cross-react with Brucella, Proteus OX19, and Yersinia. Caution: Lab should be notified of culture of possible tularemia diagnosis as needed to take precaution for work-up of cultures in BSL-3 level facility. 1 0701016F.doc 3-38 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Tularemia Table 3-7. Diagnostic Testing for Tularemia (cont.) Sputum for Gram stain, culture, DFA, and PCR Nasal/throat swab Skin lesions and biopsies: DFA and PCR and histopathology, cultures, and stains CSF for Gram stain, culture, cell count, glucose, protein, and PCR Collect > 1 mL of a lower respiratory specimen into a sterile container for Gram stain and culture. Transport in sterile, screw-capped container at room temperature when transport time is < 2 hours. For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. PCR is investigational. Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport packet at room temperature. A positive nasal swab in an asymptomatic individual following aerosol exposure only indicates exposure. A negative nasal swab does not exclude exposure. Skin lesions: Use rayon or Dacron (not cotton) swabs. Obtain a firm sample of the advancing margin of the lesion. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Aspirates: Sample should be capped (using only one hand to prevent needle stick), secured by tape, and sent to lab at room temperature (22ºC to 28°C) for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Culture media: Refer to aerobic blood cultures Biopsies: Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. Obtain 4 tubes of CSF fluid with a minimum of 2 to 3 cc per tube: Transport tube to laboratory as soon as possible at room temperature. • tube 2 – PCR Culture media: Refer to aerobic blood cultures Refrigerate (2ºC to 8ºC) if processing will be delayed. Skin biopsies: Place tissues in sterile containers. • tube 1 – cultures and stains Culture media: Refer to aerobic blood cultures Refrigerate if > 1 hour transport. • tube 3 – chemistries • tube 4 – cell count Other specimens: gastric, pleural fluid, and corneal scrapings 0701016F.doc Place tissues in sterile containers. Transport in sterile, screw-capped container at room temperature, when transport time is < 2 hours. For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. 3-39 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders GLANDERS—Bur kholder ia mallei Overview of Laboratory Exposures Background Glanders is a zoonotic disease caused by a gram-negative bacterium, B. mallei. The natural reservoirs for this organism are equine animals (horses, mules, and donkeys) and goats. Infection with this organism in horses may be systemic with prominent pulmonary involvement (known as glanders) or may be characterized by subcutaneous ulcerative lesions and lymphatic thickening with nodules (known as farcy). Although enzootic foci may exist in Asia and some eastern Mediterranean countries, glanders due to natural infection no longer occurs in the United States [1]. The method of acquisition of infection in animals is controversial but may include inhalation, ingestion, or SC inoculation. Naturally acquired glanders in humans has been generally associated with contact with infected equines. The mode of acquisition in humans is unclear but most likely occurs through various routes of exposure (e.g., inoculation through breaks in skin or nasal mucosa with infectious equine secretions or via inhalation). The infective dose in humans and animals is unknown. Humans with diabetes or renal disease are at greater risk for glanders, most likely due to impairment of white blood cells. Experience with Laboratory Exposures Laboratory-acquired infections with B. mallei have been reported to occur by inhalation and percutaneous exposures and inoculations of the skin. During the U.S. BW program (1943–1969) at Fort Detrick (then called the U.S. Army Medical Unit, Fort Detrick), 7 researchers working directly with the organism contracted glanders. Six of the infections (pulmonary glanders) occurred in the early years of the program (1943–1945) among a total of only 13 persons engaged in laboratory work with B. mallei [2]. However, all 6 individuals had significant risk of aerosolized exposure to the agent (e.g., dropping a flask containing cultures or opening lids to containers of aerated cultures in liquid medium immediately after turning off the air current) before their diagnosis of glanders. Additionally, these cases occurred during a time period when mouth pipetting and blowing out the last drop to clear the pipette (creating an aerosol) were common practices. All 6 researchers wore protective clothing, but BSCs were not yet commercially available. The incubation period in these cases was estimated to be 10 to 14 days after the suspected laboratory exposure. The researchers were diagnosed by serologies (cultures were negative) and were successfully treated for 10 to 20 days with sulfadiazine (usually 1 g every 4 hours after an initial loading dose of 4 to 5 g) [2]. Chest x-rays of these patients showed irregular densities suggestive of early lung abscesses that resolved with therapy. With changes in personal protective measures and safety education, only 1 additional case (in 1953) was diagnosed during the remainder of the BW program [3]. The only case of glanders diagnosed in the biodefense research program (1970 to present) was in March 2000. This occurred in a 33-year-old microbiologist who performed research with B. mallei at USAMRIID [4,5]. The source of exposure was thought to be from microabrasions in the skin, as the individual was known not to wear latex gloves at all times when working with the organism in the laboratory. The individual presented initially to his private physician with fever and left axillary lymphadenopathy. While symptoms improved during a 10-day course of clarithromycin, the symptoms relapsed within 4 days after discontinuation of antibiotics. The individual (known diabetic) was subsequently admitted to the hospital for diabetic ketoacidosis and multiple liver and splenic abscesses secondary to B. mallei, sensitive to imipenem, ceftazidime, and tetracycline. The individual was treated 0701016F.doc 3-40 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders with long-term antibiotics, initially IV imipenem and doxycycline for 2 weeks duration, and then oral therapy with azithromycin and doxycycline for 6 months even though drug sensitivities showed the organism resistant to azithromycin [5]. A CT scan after 6 months of antibiotic treatment showed substantial reduction in the liver and splenic abscesses, with the individual remaining in good health without recurrence 10 years later. Specific Laboratory Hazards and Biosafety Level Working Conditions B. mallei may be present in sputum, blood, wound exudates, and various tissues, depending on the site of the infection. Primary laboratory hazards are from direct contact with cultures or infectious materials, autoinoculation, inhalational exposure to infectious aerosols, and ingestion. BSL-3 facilities must be used for procedures that generate infectious aerosols and droplets with clinical materials of human or animal origin that may contain B. mallei. Animal work should be done under ABSL-3 [6]. BSL-2 facilities may be used for the primary isolations of clinical specimens and work should be done in a BSC [6]. At USAMRIID, all work with B. mallei is conducted in BSL-3/ABSL-3 containment. Experimental aerosol studies with B. mallei are conducted in a Class III BSC. The Disease Clinical Features The onset of symptoms after exposure to B. mallei is estimated to be 10 to 14 days following aerosol exposure and 1 to 5 days following percutaneous exposure. Symptoms of infection from B. mallei generally depend on the route of acquiring the infection. Infections generally manifest as (1) acute localized suppurative, (2) acute pulmonary, (3) acute septicemic, or (4) chronic suppurative forms, but the categories may overlap (Table 3-8). With early treatment, the duration of illness has ranged from 10 to 20 days. Glanders is almost always fatal without treatment. Table 3-8. Clinical Features of the Major Forms of Glanders Criterion Type of Glanders Acute Localized Acute Septicemic Suppurative Infection Infection Inoculation of abraded Inhalation or localized or lacerated skin, infection. percutaneous or mucous membrane exposure. Acute Pulmonary Infection Inhalation or hematogenous seeding. Incubation period 1 to 5 days post inoculation. 10 to 14 days. Clinical Presentation Inoculation of skin usually results in pain and swelling at the site, followed usually by a single papular or pustular lesion that gradually develops into an ulcer, associated with a mucopurulent discharge with acute lymphangitis and Route of Exposure 0701016F.doc May result in a generalized papular eruption (which may become pustular), abscess of internal organs (especially liver, spleen, lungs), and IM abscesses (especially legs and arms). Abdominal ultrasound Fever, rigors, myalgias, fatigue, headache, severe malaise, and pleuritic chest pain. Physical findings usually normal except for fever and occasional lymphadenopathy and splenomegaly. May result in extensive pneumonia. Chest x-ray may show miliary lesions, small Chronic Suppurative Infection Inoculation of abraded or lacerated skin; percutaneous or mucous membrane exposure. Multiple SC and IM abscesses located primarily on extremities (83%). 50% may have lymphatic involvement and nasal discharge or ulceration. 25% may involve pulmonary, pleural, 3-41 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders lymphadenitis (infected may show splenic or nodes may ulcerate and hepatic abscesses. drain pus). Mucous membrane involvement may cause purulent discharge from eye, nose, or lips followed by extensive ulcerating lesions; may be associated with systemic symptoms. Prognosis CFR ~20% after sulfadiazine therapy introduced. multiple lung abscesses, or ocular, skeletal, infiltrates involving upper lungs hepatic, splenic, or with consolidation and cavitation meningeal sites. but may also present as a lobar or bronchopneumonia. Note: purulent nasal discharge often seen in naturally occurring glanders was not observed in laboratory-acquired inhalational glanders. Leukocytosis may be mild; leukopenia with relative lymphoctyosis also reported. Mostly fatal in 7 to 10 days without treatment. CFR ~40% after sulfadiazine therapy introduced. Duration has ranged from 6 weeks to 15 years. CFR ~40% after sulfadiazine therapy introduced. CFR = Case Fatality Rate Chest x-rays in cases of pneumonia may demonstrate lobar or segmental opacification or diffuse nodular opacities. Cavitary lesions are common, but effusions and hilar adenopathy are rare. Abdominal ultrasound should be considered on all patients with suspected glanders to exclude the possibility of hepatic and splenic abscesses. Diagnosis Gram stain of lesion exudates may reveal small irregularly staining, gram-negative bacilli, but organisms are generally difficult to find, even in acute abscesses. Methylene blue or Wright’s stain may reveal bipolar “safety pin” staining. The organism may be cultured from abscesses, wounds, secretions, sputum (in pneumonia), and sometimes blood and urine with standard bacteriological medium; adding 1% to 5% glucose, 5% glycerol, or meat infusion nutrient agar may accelerate growth. Primary isolation requires 48 to 72 hours in agar at 37.5°C; automated blood culture methods are typically more rapid. Blood cultures for B. mallei are rarely positive upon presentation unless the patient is moribund. Identification of B. mallei by many commercial systems for rapid identification has not been reliable [7]. Serological tests do not differentiate between B. mallei and B. pseudomallei infections, due to crossreactions of the test to the agents. No validation of the serological test has been performed for diagnosis of glanders. Agglutination tests are not positive for at least 7 to 10 days (sometimes > 3 weeks), and a high background titer in normal sera (1:320 to 1:640) makes interpretation difficult. A four-fold increase in titer is considered diagnostic of glanders (a reproducible titer is recommended if a positive serological test result is obtained) [8]. Titers generally should be followed at 4 and 6 weeks after the exposure. Complement fixation (CF) tests are more specific but less sensitive and may require 40 days for conversion. CF tests are considered positive if the titer is equal to or exceeds 1:20. CF tests may not detect chronic cases of glanders. PCR tests may be useful to differentiate among B. mallei, B. pseudomallei, and B. thailandensis [9]. 0701016F.doc 3-42 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders Management of Potential Exposure to B. mallei (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to B. mallei should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure (outlined in Chapter 2) are generally followed. Diagnostic Testing • Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater-risk exposure generally require the following: – Two 10.0 mL SSTs for research serology and storage at time of exposure (should be within 7 days of exposure). – Two 10.0 mL SSTs for convalescent research serology at ≥ 14 days after incident if indicated. (Follow-up serologies also recommended at 4 and 6 weeks as delayed seroconversion may occur, especially with CF tests may require 40 days for seroconversion.) – May consider (on a case-by-case basis) nasal swabs for culture or PCR within 24 hours of inhalational exposure. A positive nasal swab within 24 hours of inhalational exposure may only indicate exposure to B. mallei, and a negative swab does not exclude exposure. Post-exposure Prophylaxis Post-exposure prophylaxis may be considered for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. There is currently no data from human studies concerning the efficacy of post-exposure chemoprophylaxis in glanders. Recommendations for chemoprophylaxis are based on results of in vitro sensitivity studies, animal studies, and success in treating the disease in humans with sulfadiazine. In vitro sensitivity testing shows that B. mallei is generally sensitive to the following parenteral antibiotics: doxycycline, macrolides, trimethoprim-sulfamethoxazole, Augmentin®, and rifampin. Most strains of B. mallei are sensitive to quinolones, but resistance has been reported to some strains [10–18]. While some strains may demonstrate resistance to macrolides, the intracellular nature of the organism and the high intracellular concentrations achieved by macrolides (cellular to extracellular ratio of > 7,000 for the drug concentration) may still render the macrolide effective, even with strains that require a high minimal inhibitory concentration (MIC) of antibiotic. Resistance to sulfanilamides has been reported [19,20]. A limited number of studies have demonstrated a protective effect with sodium sulfadiazine, doxycycline, minocycline, and quinolones in animal models. Sodium sulfadiazine was effective in the prophylaxis of glanders in hamsters when given within 24 hours of exposure and for a duration of 15 or 20 days [21]. In this study, none of 15 hamsters given 20 days of sulfadiazine and only one of 14 hamsters given 15 days of sulfadiazine relapsed, compared to relapse occurring in 18 of 30 hamsters receiving only 7 to 10 days of sulfadiazine [21]. Relapse in hamsters receiving shorter durations of therapy was observed as long as 3 to 4 weeks after cessation of antibiotic therapy (B. mallei was cultured from one or more organs). Surviving hamsters were held for 90 to 126 days; no gross pathological lesions were noted at autopsy and all cultures of organs and blood were negative. In this same study, streptomycin (initiated 3 days before challenge to achieve adequate levels in blood and tissue) only prolonged the average time of death of hamsters and was generally not effective in preventing glanders [21]. 0701016F.doc 3-43 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders Ofloxacin, doxycycline, and minocycline demonstrated activity against SC challenge in hamsters [14]. These drugs did not demonstrate appreciable activity against aerosol challenge, except for doxycycline, which provided some protection against low aerosol challenge doses of B. mallei. An abbreviated course of ciprofloxacin (5 days duration) given before or immediately after intraperitoneal challenge only delayed the course of infection, with relapse of disease occurring between days 18 to 23 of the experiment [10]. Doxycycline (5 days duration) was superior to ciprofloxacin in protecting hamsters. No relapses occurred before Day 23 of the experiment, but relapses were observed at 4 to 5 weeks in 4 of 8 hamsters that began treatment on Day 0 of challenge [10]. Similarly, studies in BALB/c mice given delayed therapy (initiated 24 hours after intranasal challenge) with levofloxacin or ceftazidime (10 days duration) resulted in 100% survival rate at Day 34 but did not eradicate disease [15]. While viral burdens were greatly reduced in treated mice compared to untreated controls, chronic multifocal abscesses were observed in the spleen (bacterial burdens were lower with levofloxacin compared to ceftazidime). A more recent study demonstrated various quinolones, if initiated within 3 hours after inoculation and for 10 days duration, were 80% to 90% effective in preventing disease compared to only 60% to 90% protection with a 15-day course of quinolones that was initiated at 24 hours following exposure (follow-up for 30 days after cessation of antibiotics) [17]. As hamsters are more susceptible to infection than humans, the risk of relapse in humans may be lower. Parenteral antibiotics that have demonstrated in vitro sensitivity to B. mallei are summarized in Table 3-9. Table 3-9. Parenteral Antibiotics Demonstrating Sensitivity on in vitro Sensitivity Tests to B. mallei Drug Trimethoprim–Sulfamethoxazole DS Doxycycline Tetracycline Augmentin Macrolides Quinolones Comment 1 tablet 2 times daily. Sulfadiazine was protective in hamsters without relapse if given for 20 days and within 24 hours of exposure [21] 100 mg 2 times daily. B. mallei highly sensitive to doxycycline on in vitro sensitivity tests; greater protection than quinolones in hamsters. Auromycin was successful in treating a lone case of glanders. 500 mg 4 times daily 500 mg tablet 3 times daily Azithromycin, clarithromycin, erythromycin. High intracellular concentrations with macrolides (cellular to extracellular ratio of > 7000 for drug concentration) may still render a macrolide effective even with strains that require a high MIC of antibiotic. Ciprofloxacin 500 mg 2 times daily. Some resistance reported in vitro, but quinolones have high intracellular concentrations; one animal study shows effectiveness as postexposure chemoprophylaxis if given within 3 hrs of exposure but associated with relapse of disease in other studies when initiated ≥ 24 hrs post-exposure [10,15, 17]. Males and Non-pregnant Females If chemoprophlyaxis is recommended, it should be started as soon as possible (preferably within 24 hours after exposure, based on animal studies). Modifications of chemoprophylaxis regimens should be based on results of sensitivity testing, if available. There currently are no data concerning the use of one versus two drugs (as required in treatment regimens) for post-exposure prophylaxis of glanders in humans or data concerning the duration of postexposure prophylaxis therapy. Based on sulfadiazine (20 days) being successful in treating humans with glanders and protecting hamsters post challenge with B. mallei, trimethoprim-sulfamethoxazole may be considered a drug of 0701016F.doc 3-44 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders choice if post-exposure prophylaxis is recommended (if the organism is not known to be resistant to trimethoprim-sulfamethoxazole). The duration of chemoprophylaxis in humans is unknown. Based on the relapses occurring in hamsters given ≤ 10 days of sulfadiazine but not with 20 days of sulfadiazine, one may consider 21 days of chemoprophylaxis. If trimethoprim-sulfamethoxazole cannot be used, one may consider using doxycycline alone or combined with a second drug (as required for treatment of glanders), realizing there currently are no data concerning an increased benefit from the use of two drugs for prophylaxis compared to doxycycline alone in humans. Individuals need to be closely monitored after cessation of prophylaxis for relapse of disease. Serological testing should be obtained for at least 6 weeks after the exposure. If serological testing demonstrates seroconversion, the individual should receive a treatment course of antibiotics for glanders (see “Treatment of Overt or Suspected Glanders”). Pregnant Females Recommendations for antibiotic prophylaxis in pregnant women with significant risk exposure to B. mallei: • Consultation with infectious diseases and/or obstetrics experts (or other experienced health care provider) is highly recommended. Post-exposure prophylaxis must be assessed on an individual basis. The risk of post-exposure prophylaxis must be weighed against the exposure and risk of disease from B. mallei. • B. mallei strains are generally sensitive to amoxicillin-clavulanate (Augmentin), which is rated as Category B for use in pregnancy. Macrolides may also be considered. Ciprofloxacin is a Pregnancy Category C drug. Although the TERIS concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Doxycycline is a Category D agent in pregnancy that may cause discoloration of fetus teeth if given during the last half of pregnancy. Management of Suspected Glanders (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to B. mallei must be recorded with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Blood for culture • Whole blood in citrate tubes for PCR (may use serum or citrated plasma as secondary source) • Serum for immunoassays (acute and convalescent sera) • Sputum, if present, for Gram stain, culture, and PCR • Pathology specimens for stain, culture, histopathology, and PCR • Any purulent drainage, if present, for Gram stain and culture Collection and Handling of Clinical Specimens Blood Cultures Collect appropriate blood volume and number of sets of blood cultures (usually 2 sets initially) and then as clinically indicated. BACTEC adult culture bottles require 10 mL per bottle. If the patient has 0701016F.doc 3-45 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders received antibiotics, blood cultures containing resin to absorb the antibiotics are recommended. Transport samples directly to the laboratory at room temperature. Hold them at room temperature until they are placed on the blood culture instrument or incubator. Do not refrigerate. Serum Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing; transport chilled and not frozen. Collect one 7.0 mL red top or TT tube for serum for serologies (may refrigerate at 2ºC to 4ºC for 5– 7 days; freeze at -30ºC or lower if > 7 days). Respiratory Cultures Sputum and induced respiratory secretion specimens for culture and PCR should be placed in sterile, screw-capped containers at room temperature. Material to be transported from 2 to 24 hours after collection should be stored in transport container at 2ºC to 8ºC. Swabs (rayon or Dacron only, never cotton) should be moistened with the medium inside the packet and reinserted into the transport package. Specimens should be transported at room or refrigerator (2ºC to 8ºC) temperature. Nasal Swabs Use rayon or Dacron (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Skin Lesions Biopsied tissue or scraping of an ulcer is preferable, but a swab of an ulcer is an acceptable alternative. Submit tissue or scraping in a sterile container. Transport at room temperature for immediate processing at USAMRIID. If processing of specimen is delayed, specimen should be kept chilled (2ºC to 8ºC). Biopsies Collect biopsy, scraping, swab, or aspirate of ulcer, lymph node, or other involved tissue. For biopsy, place the tissue or scraping in a sterile container. Transport at room temperature for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Treatment of Confirmed or Suspected Glanders The limited number of laboratory infections in humans has precluded evaluation of effectiveness of most antibiotics. Sulfadiazine has been effective in treatment of glanders in humans [2]. Animal experiments indicate that sulfadiazine, ceftazidine, imipenem, tetracyclines, and quinolones may be effective in the treatment of glanders. In vitro studies have demonstrated that most strains of B. mallei are sensitive to ceftazidime, imipenem, piperacillin, Unasyn, doxycycline, rifampin, trimethoprim-sulfamethoxazole, and macrolides [11,12]. Although the organism is sensitive to piperacillin, piperacillin-tazobactam should be considered instead as it has a three- to fourfold lower minimal inhibitory concentration than piperacillin. Most (but not all) strains of B. mallei are sensitive to quinolones. Increased resistance to quinolones has been reported [9– 11,13,15–17], as well as an increase in resistance to co-trimoxazole [18]. Antibiotics having higher intracellular concentrations (e.g., macrolides and quinolones) may have a higher therapeutic effect than 0701016F.doc 3-46 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders that demonstrated by in vitro sensitivity testing. Macrolides have a cellular to extracellular ratio of > 7,000 of drug concentration and may still be effective even though some strains demonstrate resistance. Although B. mallei may demonstrate in vitro sensitivity to aminoglycosides, the poor intracellular penetration of aminoglycosides suggests that this class of drugs may not be as therapeutically effective. Treatment of glanders involves both an intensive and an eradication phase of therapy, similar to the treatment regimen recommended for B. pseudomallei infection (melioidosis). Intensive Therapy IV therapy is recommended initially for a minimum period of 10 to 14 days (longer treatments may be required for severely ill patients or deep-seated locations), generally initiated in conjunction with prolonged eradication therapy (given orally). Options for initial parenteral treatment include imipenem (25 mg/kg, up to 1 g) every 6 hours, meropenem (25 mg/kg, up to 1 g) every 8 hours, or ceftazidime (50 mg/kg, up to 2 g) every 6 hours combined with or without trimethoprim-sulfamethoxazole (8/40 mg/kg, up to 320mg/1600 mg) every 12 hours. Trimethoprim-sulfamethoxazole is recommended to be included in the treatment regimen for neurologic, cutaneous, bone, and prostate infections. Eradication Therapy Oral eradication therapy is generally initiated concomitantly with the intensive IV therapy [10–12]. Duration of oral therapy may depend on the extent and severity of disease. Based on the experience in treatment of melioidosis, oral therapy for 12 to 20 weeks duration may be recommended for disease localized to the lungs, but prolonged therapy (6 to 12 months) may be required for deep-seated infections (i.e., severe disease and visceral disease, CNS infections, prostate infections). Trimethoprimsulfamethoxazole, either alone or in combination with doxycycline, is generally the initial recommended treatment of choice for eradication therapy unless the organism is known to be resistant to trimethoprimsulfamethoxazole. Alternate therapy (if trimethoprim-sulfamethoxazole cannot be given) may include the following antibiotics (a combination of two drugs may be recommended): doxycycline, macrolides, chloramphenicol, quinolones, or amoxicillin-clavulanate. Drug therapy experience in patients with melioidosis in humans has demonstrated (1) an increased risk of relapse with doxycycline alone for eradication therapy (doxycycline monotherapy thereby not recommended as first-line eradication therapy) and (2) an increased risk of relapse with amoxicillin-clavulanate and quinolones compared to trimethoprim-sulfamethoxazole, doxycycline, and chloramphenicol for eradication therapy [7]. Drugs for Eradication Therapy of Glanders Trimethoprim-sulfamethoxazole (8/40 mg/kg, up to 320/1600 mg) may be given every 12 hours (generally for 60 to 150 days) alone or in combination with doxycycline (2.5 mg/kg, up to 100 mg) every 12 hours or chloramphenicol (generally first 4 weeks only when used in triple drug therapy). Alternatives include the macrolides, amoxicillin-clavulanate (adult dosage of 500 mg every 8 hours or 875 mg every 12 hours), or quinolones [7]. Sensitivity testing should be performed to aid with therapy. Surgical Drainage of Abscesses Surgical drainage depends on the location and should be performed when possible. Follow-up Patients require follow-up at regular intervals for at least 5 years and may require lifelong follow-up. Patients should be alerted to the life-long risk of relapse and they need to alert healthcare providers of a 0701016F.doc 3-47 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders history of glanders, particularly when being evaluated for febrile illnesses or other symptomatology that could represent a relapse of glanders. Environmental Decontamination and Infection Control B. mallei is not hardy and cannot survive in dried pus for longer than a few days or survive exposure to sunlight for longer than 24 hours. The organism is also killed by exposure to moist heat at 55°C for 10 minutes and also by disinfectants (e.g., 1% sodium hypochlorite). However, the organism may remain viable in tap water for 1 month or longer. Person-to-person transmission of B. mallei can occur and is the second most common source of natural disease in humans next to acquisition from infected horses. Private room placement and droplet precautions (mask if within 3 to 6 feet of a patient that has B. mallei pneumonia or during procedures such as suctioning) are required. References 1. Heymann DL, ed: Control of Communicable Diseases Manual, 18th ed, p 356. Washington DC, American Public Health Association, 2004. 2. Howe C, Miller WR: Human glanders: report of six cases, Ann Intern Med 1947; 26(1):93–115. 3. Rusnak JM, Kortepeter MG, Hawley RJ, Anderson AO, Boudreau E, Eitzen E: Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers, Biosecur Bioterr 2004; 2(4):281–93. 4. DeShazer D, Byrne WR, Culpepper R, Andrews G, Hartman L, Parker G, Heine H, Belani A, boyer J, Barrera-Oro M, Kraus C, Srinivasan a, Karenfil L, Perl T, Bartlett J, Bowes J, Smith J, Denner A, Hankinson A, Edwards L, Roche J, Laboratory-acquired human glanders—Maryland, May 2000, MMWR Morb Mortal Wkly Rpt 2000; 49(24):532–35. 5. Srinivasan A, Kraus CN, DeShazer D, Becker PM, Dick JG, Byrne WR, Thomas DL: Glanders in a military research microbiologist, N Engl J Med 2001; 345(4):256–58. 6. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, and National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed, pp 126–28. Washington DC, U.S. Government Printing Office, 2007. www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5TOC.htm. Accessed December 7, 2007. 7. Gilad J: Burkholderia mallei and Burkholderia pseudomallei: the causative micro-organisms of glanders and meliodosis, Recent Pat Antiinfect Drug Discov 2007; 2(3):233–41. 8. Peacock SJ, Schweizer HP, Dance DAB, Smith TL, Gee JE, Wuthiekanun V, DeShazer D,Steinmetz I, Tan P, Currie BJ: Management of accidental laboratory exposure to Burkholderia pseudomallei and B. mallei [Online report], Emerg Infect Dis 2008; 14(7):e2. 9. Tanpiboonsak S, Paemanee A, Bunyarataphan S, Tungpradabkul S: PCR-RFLP based differentiation of Burkholderia mallei and Burkholderia pseudomallei, Mol Cell Probes 2004; 18(2):97–101. 10. Russell P, Eley SM, Ellis J, Green M, Bell DL, Kenny DJ, Titball RW: Comparison of efficacy of ciprofloxacin and doxycycline against experimental melioidosis and glanders, J Antimicrob Chemother 2000; 45:813–18. 11. Heine HS, England MJ, Waag DW, Byrne WR: In vitro antibiotic susceptibilities of Burkholderia mallei (causative agent of glanders) determined by broth microdilution and E-test, Antimicrob Agents Chemother 2001; 45(7):2119–21. 12. Thibault FM, Hernandez E, Vida DR, Girardet M, Cavallo J-D: Antibiotic susceptibility of 65 isolates of Burkholderia pseudomallei and Burkholderia mallei to 35 antimicrobial agents, J Antimicrob Chemother, 2004; 54(6):1134–38. 0701016F.doc 3-48 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders 13. Stepanshin IuG, Manzeniuk IN, Svetoch EA, Volkovoĭ KI: In vitro development of fluoroquinolone resistance in the glanders pathogen [Russian], Antibiot Khimioter 1994; 39(8):30–33. 14. Iliukhin VI, Alekseev VV, Antonov IuV, Savchenko ST, Lozovaia NA: Effectiveness of treatment of experimental glanders after aerogenic infection [Russian], Antibiot Khimioter 1994; 39(9–10):45–48. 15. Judy BM, Whitlock GC, Torres AG, Estes DM: Comparison of the in vitro and in vivo susceptibilities of Burkholderia mallei to ceftazidime and levofloxacin, BMC Microbiol 2009; 9:88. 16. Bondareva TA, Kalininskiĭ VB, Borisevich IV, bondarev VP, Kozhukhov VV, Amosov Mlu, Baramzina GV, Fomenkov OO, Krinitsyn FV: Fluroquiniolones: antimicrobial activity and chemotherapeutic efficacy with respect to various pathogens in highly dangerous diseases [Russian], Antibiot Khimioter 2007; 52(11–12):21–24. 17. Bondareva TA, Borisevich IV, Kalininskiĭ VB, Vondarev VP, Fomenkov OO: Study of efficacy of modern fluoroquinolones against agent of glanders in experiments in vivo [Russian], Zh Microbiol Epidemiol Immunobiol 2009 May–Jun; 3:10–13. 18. Kenny DJ, Russell P, Rogers D, Eley SM, Titball RW: In vitro susceptibilities of Burkholderia mallei in comparison to those of other pathogenic Burkholderia spp, Antimicrob Agents Chemother 1999; 43(11):2773–75. 19. Manzeniuk IN, Stepanshin IuG, Svetoch EA: Evaluation of the potential of chemotherapy of glanders caused by Pseudomonas mallei strains resistant to sulfanilamides and trimethoprim, Antibiot Khimioter 1994; 39(7):30–34. 20. Batmanov VP, Iliukhin VI, Lozovaia NA, Iakovlev AT: Recovery rate in chemotherapy of glanders [Russian], Antibiot Khimioter 1996; 41(5):30–34. 21. Miller WR, Pannell L, Ingalls MS: Experimental chemotherapy in glanders and melioidosis, Am J Hyg 1948; 47(2):205–13. 0701016F.doc 3-49 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders Table 3-10. Diagnostic Testing for Glanders Specimen Site and Test Aerobic blood cultures Serum for PCR Method of Collection Transport Comments Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Transport at room temperature. Whole blood is preferable (collected in citrate tube) for PCR; serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Transport chilled (not frozen). Store at 4ºC (not frozen). PCR tests, though not validated, may be useful in differentiating among B. mallei, B. pseudomallei, and B. thailandensis. Refrigerate at 2ºC to 4ºC for 5 to 7 days. Agglutination tests: Not positive for at least 7 to 10 days; high background titer in normal sera (1:320 to 1:640) makes interpretation difficult. Do not refrigerate. Further sets of blood cultures obtained as clinically indicated. Blood cultures are rarely positive upon presentation unless patient is moribund. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for PCR. Collect 1 mL minimum (7 mL adequate) of serum for serology in either red top or TT tubes and spun. Freeze at -30ºC or lower if greater than 7 days. Fourfold increase in serology is diagnostic of infection. A single agglutination titer > 1:160 with compatible clinical picture suggests active infection of melioidosis or glanders. Serum for acute and convalescent serologies CF tests more specific but less sensitive and may require 40 days for seroconversion (fourfold increase in titer). Often considered positive if titer ≥1:20. CF tests may not detect chronic cases of glanders. Sputum for Gram stain, culture, and PCR Nasal/throat swab 0701016F.doc Collect >1 mL of a lower respiratory specimen into a sterile container for stains and culture. Transport in sterile, screw-capped container at room temperature, when transport time is <2 hours. For transport times between 2 to 24 hours, transport at 2º to 8ºC. Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport packet at room temperature. Gram stain small irregularly staining gram-negative bacilli. Methylene blue or Wright stain may reveal bipolar “safety pin” appearance. Organisms generally difficult to find, even in acute abscesses. Refrigerate at 2ºC to 8ºC if processing is delayed. 3-50 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Glanders Skin lesions and biopsies: PCR, histopathology cultures, and stains 0701016F.doc Skin lesions: Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Skin biopsies: Place tissues in sterile containers. Aspirates: Sample should be capped, secured by tape, and sent to lab at room temperature (22ºC to 28°C) for immediate processing. Refrigerate (2ºC to 8ºC) if processing will be delayed. Biopsies: Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. 3-51 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis MELIOIDOSIS—Bur kholderia pseudomallei Overview of Laboratory Exposures Background Melioidosis is a zoonotic disease caused by a gram-negative bacterium, B. pseudomallei. The organism is saprophytic in soil and water and is endemic in Southeast Asia, Northern Australia, China, and India. Sporadic cases are reported from Central and South America, Africa, the Caribbean, and the Middle East. Various animals, including sheep, goats, horses, swine, monkeys, and rodents (along with a variety of animals and birds in zoological gardens) can become infected. B. pseudomallei can be spread to humans through the nasal, oral, and conjunctival mucous membranes and by inhalation into the lungs, but it is thought to be most predominantly acquired through direct contact of skin abrasions (with contaminated soil or water). Vertical transmission at childbirth and sexual transmission have been reported but are uncommon. In endemic areas, 5%–20% of agricultural workers may have antibody titers to B. pseudomallei without overt disease [1]. Risk factors for development of melioidosis are diabetes mellitus, chronic renal or lung disease, and excess alcohol consumption. Experience with Laboratory Exposures Infection of laboratory workers with B. pseudomallei has been rare [2,3]. Only two laboratory-associated cases of melioidosis (neither at USAMRIID) have been reported [4,5]. One occurred when a centrifuge developed a leak and a suspension of the organism was sprayed over the walls and work benches. A worker cleaned the area with her bare hands, putting her at risk for both aerosol exposure from the break in the centrifuge and direct exposure to the agent while cleaning up. As a result, she developed an ulcerative lesion at the base of her right index finger and pneumonia. The second case occurred when an aerosol was created during open-flask sonication of B. pseudomallei outside the BSC (originally misidentified as Pseudomonas cepacia) [4,5]. In a study published in 1992, serologic follow-up of 60 laboratory workers from a clinical laboratory in a melioidosis-endemic area over a 15-year period indicated the organism poses a low risk for laboratory workers following a laboratory policy to minimize exposure. In this laboratory setting, all aerosolgenerating procedures were performed in a Class II or higher BSC, whereas new cultures were examined on the open bench with sniffing of cultures prohibited. When tested, only 3 of these 60 laboratory workers had elevated B. pseudomallei titers (suggesting a past subclinical infection), which was consistent with the seroprevalence in the community [6]. In 2003, 13 laboratory workers were identified as having a high risk for exposure to B. pseudomallei from sniffing culture plates and/or performing routine laboratory procedures such as subculturing and inoculating the B. pseudomallei outside a BSC (before organism was identified). These individuals were given post-exposure prophylaxis with a 2-week course of trimethoprim-sulfamethoxazole beginning Day 0 to Day 4. Based on results from follow-up serologies at 6 weeks, none of the individuals became ill or developed antibodies to B. pseudomallei, which would indicate either a low risk of infection for laboratory workers and/or possibly the effectiveness of post-exposure prophylaxis [7]. In 2005, 9 laboratory workers were identified as having risk of exposure to B. pseudomallei from handling the organism outside a BSC. Only the 3 individuals who sniffed the culture plates received post-exposure prophylaxis, even though post-exposure prophylaxis was offered to all 9 individuals. No individuals developed symptoms of melioidosis or seroconverted [2]. 0701016F.doc 3-52 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis Specific Laboratory Hazards and Biosafety Level Working Conditions B. pseudomallei has been demonstrated in blood, sputum, wound exudates, and various tissues, depending on the site of infection [1], and may be present in soil and water samples from endemic areas. B. pseudomallei may persist in the environment for years in moist soil or water, and as long as 4 weeks in dry soils. Direct contact with cultures and infectious materials from humans, animals, or the environment; ingestion (uncommon); autoinoculation; and exposure to infectious aerosols and droplets are the primary laboratory hazards of working with B. pseudomallei. BSL-3 facilities must be used for procedures that generate infectious aerosols and droplets with clinical materials of human or animal origin that may contain B. pseudomallei. BSL-2 facilities may be used for clinical specimens and cultures (provided work is not associated with production of aerosols or droplets), with work performed in a BSC [1,8]. Animal studies should be conducted under ABSL-3 [1]. At USAMRIID, all work with B. pseudomallei is conducted in BSL-3/ABSL-3 containment. Experimental aerosol studies with B. pseudomallei are conducted in a Class III BSC. The Disease Clinical Features The melioidosis incubation period varies by route of entry, size of inoculum, virulence of organism, and host factors. Most individuals exposed to B. pseudomallei do not develop symptoms. Those who do develop symptoms usually have predisposing medical conditions (e.g., diabetes mellitus, chronic renal failure, chronic obstructive airway disease, daily excess alcohol consumption, or immunosuppression from steroids). Individuals with cystic fibrosis or bronchiectasis may have colonization of their airways with B. pseudomallei (and subsequent disease) and should be cautioned about the risk of disease [9]. The organism may remain latent for a number of years (decades) after initial acquisition and before symptoms from disease occur, often in association with the onset of an immunocompromising state. Duration of illness is uncertain due to the variable nature of the disease. The type of infection and the course of treatment can predict long-term sequelae. Death is rare in patients who do not have predisposing risk factors and is generally only associated with B. pseudomallei bloodstream infection. Even after prolonged antimicrobial therapy, relapses are common. While there are different classifications for melioidosis, infections generally manifest as (1) acute pulmonary, (2) acute localized, (3) acute septicemic, and (4) chronic suppurative, but the categories may overlap (Table 3-11). Acute pulmonary infection is the most common form of melioidosis and primary pneumonia is the most common clinical presentation, accounting for approximately half of the melioidosis cases. Secondary pneumonia accounts for approximately 10% of melioidosis cases (seeding from primary sites). Melioidosis may also present as a mild undifferentiated pneumonia (subacute or acute infection) with low mortality or as septic shock with a mortality of 90%. Chronic infection (> 2 months duration) may involve the lungs and may present similar to tuberculosis (fever, weight loss, cough, hemoptysis, and infiltrates or cavitary lesions of the upper lobes of the lung). Acute localized B. pseudomallei infection may present as an ulcer, abscess, or cellulitis [10,11] associated with acute lymphangitis, which may progress to acute septicemic form if not treated. The acute septicemic form of disease is generally acquired from inhalation or secondary to a localized infection. Bacteremia has been reported in 60% of melioidosis cases in Thailand and 46% of cases in Australia, with mortality rates of 44% and 19%, respectively, in the two countries [11,12]. Approximately 20% of melioidosis cases developed septic shock (associated with a 86% mortality rate) [11]. Most deaths from 0701016F.doc 3-53 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis melioidosis were generally associated with septic shock, with only 6 deaths reported out of 202 cases without septic shock in a large cohort in Austrailia [11]. Mortality is generally lower for localized disease without bacteremia (< 10%). Mortality is higher for septicemic melioidosis if associated with bacteremia and one organ system involvement (10% to 40%) or with bacteremia and multiple organ system involvement (40% to 50%), or with septic shock (80% to 90%) [13]. Table 3-11. Clinical Features of the Major Forms of Melioidosis Types of Melioidosis Criterion Acute Pulmonary Infection Inhalation or secondary from septicemia. Route of Exposure Incubation Period Inhalation or secondary from localized infection. Chronic Suppurative Infection Inoculation of skin or percutaneous or mucocutaneous exposure. Variable, but generally 1–21 days with an average of 9–14 days; May have asymptomatic exposure with reactivation of disease years later. 1 to 5 days postinoculation. Variable, but generally 1 to 21 days [14]. Variable, but generally 1 to 21 days [1]. Most common form of the disease. Inoculation of skin usually results in ulcer or abscess or cellulitis at site of inoculation with acute lymphangitis. Fever, headache, pharyngitis, diarrhea, cutaneous pustular lesions (uncommon), severe muscle tenderness, and pneumonitis. Multiple SC and IM abscesses located primarily on extremities (83%). Primary pneumonia or secondary pneumonia from hematogenous seeding; May present as mild undifferentiated pneumonia (onset abrupt or subacute) or severe pneumonia with sepsis. Chest pain, cough, hemoptysis, mild pharyngitis, and tachypnea. Chest x-ray may have lobar pneumonia, diffuse nodular infiltrates that later cavitate. NOTE: Chronic presentation (lasting over 2 months) may present similar to tuberculosis with fever, 0701016F.doc Inoculation of skin or percutaneous or mucocutaneous exposures. Acute Septicemic Infection Chronic pulmonary infection. Fever ≥ 102°F. Clinical Presentation Acute Localized Infection Fever and malaise. May progress to acute septicemic form. Relapse common even after prolonged antimicrobial therapy. Hepatosplenomegaly may ensue with cyanosis. Onset may be dramatic with presentation of shock. Organs involved include skin, brain, lung, myocardium, liver, spleen, bones, joints, lymph nodes, and eye. Patients may remain afebrile, even in presence of multiple abscesses. Chronic presentation in lungs (lasting over 2 months) may present similar to tuberculosis with fever, weight loss, hemoptysis, and cavitary lesions in upper lungs. 3-54 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis weight loss, hemoptysis, and infiltrates or cavitary lesions in upper lobes of the lungs. Prognosis Variable. Lower mortality if no bacteremia (< 10%). Higher mortality if (1) sepsis or bacteremia and one organ system involvement (10% to 40%) or multiple organ system involvement (40% to 50%) or (2) septic shock (80% to 90%). Localized disease involving a single organ system without bacteremia usually associated with mortality rate of < 10%. 80% to 90% fatality within 24 to 48 hours of onset of septic shock. Death generally uncommon in chronic meliodosis in absence of predisposing risk factors or sepsis. Mortality (treated) for severe melioidosis is 44% in Thailand; 19% in Australia. Differential diagnosis of melioidosis includes glanders, which is caused by B. mallei. Local nodule/abscess formation and regional lymphadenitis are more commonly seen in glanders than in melioidosis patients. Chronic pulmonary infection (lasting over 2 months) may present similar to tuberculosis with fever, weight loss, hemoptysis, and infiltrates or cavitary lesions in upper lobes of the lungs. Diagnosis A detailed summary of the diagnosis of melioidosis is presented in two published reviews [15,16]. Methylene blue or Wright’s stain of exudates may reveal scant small bacilli with a safety-pin bipolar appearance. However, the morphology of the organism may be variable in clinical specimens. Isolation of B. pseudomallei from body fluids remains the “gold standard” for diagnosis of melioidosis. Standard cultures can be used to identify both B. mallei and B. pseudomallei, but selective media (modified Ashdown’s medium) have been used for isolation of B. pseudomallei from nonsterile sites such as sputum and pharyngeal cultures [17]. The addition of 1%–5% glucose, 5% glycerol, or meat infusion nutrient agar may accelerate growth. Primary isolation of B. mallei requires 48–72 hours in agar at 37.5°C; automated blood culture methods are typically more rapid. B. pseudomallei is generally faster growing and less fastidious than B. mallei. Commercial identification systems may vary in ability to diagnose B. pseudomallei [13,17], and many commercial systems for rapid identification are not reliable for identification of B. pseudomallei (new software release with version 4.03 for VITEK 2 60/XL may have improvements for identification of B. pseudomallei) [13,18]. Blood cultures for B. pseudomallei septicemia are often positive, with the organism detected within 48 hours in 90% of positive blood cultures. Sputum culture should be obtained in pulmonary infections. If a sputum specimen is not available, throat cultures (with swabs) have a sensitivity of 36% using selective media (sensitivity of 24% if sputum culture is positive and 79% if sputum culture is negative for B. pseudomallei) [19]. Urine culture may be positive, especially if prostatitis or renal abscesses are present [15]. 0701016F.doc 3-55 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis Serological tests for melioidosis may cross-react with glanders. Serological tests do not differentiate between B. pseudomallei and B. mallei infections. Agglutination tests are not positive for at least 7–10 days. A high background titer in normal sera (1:320 to 1:640) may make interpretation difficult. For B. pseudomallei, a fourfold increase in titer supports the diagnosis of melioidosis (a reproducible fourfold increase in titer is recommended if a positive titer occurs after exposure) [8]. A single titer > 1:160 with a compatible clinical picture suggests active infection for either B. mallei or B. pseudomallei. For individuals from nonendemic areas, an agglutination titer of ≥ 1:40 with a compatible clinical picture or a known negative baseline titer may be significant [8,13]. CF tests are more specific and are considered positive if a titer is ≥ 1:20. Titers are recommended for 4–6 weeks following exposure as delayed seroconversion has been reported, particularly with CF tests. PCR tests may be useful to differentiate among B. pseudomallei, B. mallei, and B. thailandensis [20]. Bacterial fatty acid methyl ester (FAME) profile analysis by gas-liquid chromatography (GLC) to detect a cellular fatty acid profile may be used to distinguish B. pseudomallei from B. thailandensis [21]. Latex agglutination tests (based on polyclonal or monoclonal antibodies to lipopolysaccharide or exopolysaccharide) are reported to be more specific and are used for diagnosis of melioidosis in Thailand but are not commercially available [14,22–24]. Direct immunofluorescent microscopy techniques (including rapid tests that may be performed within 10 minutes) for diagnosis of B. pesudomallei in clinical samples (pus, respiratory secretions, and urine) have also been used (66% sensitivity and > 99% specificity compared to culture) [25]. Management of Potential Exposure to B. pseudomallei (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to B. pseudomallei should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure (outlined in Chapter 2) are generally followed. Diagnostic Testing • Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater-risk exposure generally require the following: – Two 10.0 mL SSTs for research serology and storage at time of exposure (should be within 7 days of exposure). – Two 10.0 mL SSTs for convalescent research serology at ≥ 14 days post-exposure (follow-up serologies are also recommended at 4–6 weeks following the exposure), if indicated, for serologic diagnosis. – May consider (on a case-by-case basis) nasal swabs for culture or PCR within 24 hours of inhalational exposure. A positive nasal swab within 24 hours of an inhalational exposure incident only indicates exposure to B. pseudomallei, and a negative swab does not exclude exposure. Post-exposure Prophylaxis Post-exposure prophylaxis may be recommended for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. There are currently no data from human studies concerning the efficacy of post-exposure chemoprophylaxis in melioidosis. The 0701016F.doc 3-56 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis recommendation for chemoprophylaxis is based on results of in vitro sensitivity studies and a limited number of animal studies [12, 26-29]. Oral agents that have demonstrated efficacy on in vitro testing include trimethoprim-sulfamethoxazole, doxycycline, tetracycline, chloramphenicol, and Augmentin (amoxicillin-clavulanate potassium). Some strains are sensitive to quinolones, but as high as 50% of strains demonstrated resistance or intermediate sensitivity on in vitro tests. B. pseudomallei is intrinsically resistant to macrolides and aminoglycosides. Sodium sulfadiazine was effective in prophylaxis of melioidosis in hamsters (100% survival) if given within 6 hours of exposure whether given for a duration of 10 days (N = 7) or 20 days (N = 8), without evidence of disease on autopsy performed after a 90–110 day observation period [26]. Only 50% survival (4 of 8 animals) was observed when initiation of sodium sulfadiazine therapy was delayed to 21 hours post-inoculation and given for 10 days duration. Hamsters exhibited relapse of disease after therapy was discontinued. More recently published studies in animals demonstrated chemoprophylaxis was most effective if given before or immediately following challenge with B. pseudomallei and generally was much less effective if initiation of chemoprophylaxis was delayed to 24 hours or longer after challenge [27-29]. One animal study compared the efficacy of doxycycline, amoxicillin-clavulanic acid, and co-trimoxazole for postexposure prophylaxis in BALB/c mice against aerosol exposure to B. pseudomallei [29]. Co-trimoxazole given for 10 days was found to be superior, with a 100% survival rate when initiated at 48 hours before aerosol challenge and when initiated at 0, 10, or 24 hours after challenge with B. pseudomallei, but not when initiated at 48 hours following challenge. Doxycycline was slightly less effective. Survival rates at Day 21 post-exposure were 100%, 80%, 60%, and 0% when started at 0 hours, 10 hours, 24 hours, and 48 hours post-challenge, respectively. Augmentin was not effective as post-exposure prophylaxis, with 100% mortality of all mice in all 5 groups by day 8 post-exposure (5 mice/group). Post-mortem exam at Day 21 of surviving mice that were given either co-trimoxazole or doxycycline was negative for culture of B. pseudomallei from the lungs, liver, or spleen (post-mortem exam of mice with relapse revealed spenomegaly and multiple small abscesses in the lungs and liver that were not as severe as those in the control mice). In another study, post-exposure prophylaxis in white rats with quinolones or trimethoprimsulfamethoxazole (10 days duration of drug) initiated within 3 hours of subcutaneous exposure was 100% effective in preventing the disease (verified by necropsy at 2 months post-exposure) [28]. Another animal study demonstrated early administration (48 hours before challenge or immediately after challenge) of either doxycycline or ciprofloxacin for 5 days duration was protective in most mice against challenge with B. pseudomallei [27]. However, a few animals had relapse of infection during the 5 week observation period. Post-mortem exam of surviving animals revealed splenomegaly and abscesses in some mice and also positive cultures for B. pseudomallei in some mice (even without evidence of gross infection). If initiation of antibiotics was delayed to 24 hours post-exposure, relapses (usually manifested by splenic abscesses) were observed in most mice (particularly between days 11 and 21 after challenge). Males and Non-pregnant Females There is currently no controlled trial in humans concerning the efficacy of postexposure prophylaxis in melioidosis. If prophylaxis is recommended, it should be given as soon as possible (within a few hours of the exposure, based on animal studies). A 21 day treatment regimen may be considered, particularly if chemoprophylaxis was delayed ≥ 24 hours following the exposure (a 14 day regimen was also highly effective in most animals and may be considered if individuals cannot complete a 21 day course). There 0701016F.doc 3-57 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis currently are no data concerning the use of one versus two drugs (as required in treatment) for postexposure prophylaxis for melioidosis. The relapse in animals with monotherapy suggests perhaps a combination of two drugs and a duration of at least 14–21 days be administered, particularly if postexposure prophylaxis was delayed more than a few hours. Modifications of chemoprophylaxis regimens should be based on results of sensitivity testing, if available. Based on animal studies, one may consider chemoprophylaxis with trimethoprim-sulfamethoxazole (1 double-strength tablet 2 times daily). If trimethoprim-sulfamethoxazole cannot be given, the alternate regimen should include doxycycline (100 mg every 12 hours), if possible. Other alternative or additional oral drugs for post-exposure prophylaxis include tetracycline (in place of doxycycline), chloramphenicol, and possibly Augmentin (amoxicillin-clavulanate potassium) or quiniolones (if known to be sensitive to quinolones). Individuals should be closely monitored for relapse after chemoprophylaxis is discontinued. Pregnant Females Recommendations for antibiotic prophylaxis in pregnant women with significant risk of exposure to B. pseudomallei: • Consultation with infectious diseases and/or obstetrics experts (or other experienced health care provider) is highly recommended. Post-exposure prophylaxis must be assessed on an individual basis. The risk of post-exposure prophylaxis must be weighed against the exposure and risk of disease from B. pseudomallei. • B. pseudomallei strains are generally sensitive to amoxicillin-clavulanate (MIC 1-4), which is rated as Pregnancy Category B drug. Ciprofloxacin is a Pregnancy Category C drug. Although the TERIS concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Management of Suspected Melioidosis (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to B. pseudomallei must be documented with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Blood for culture. • Whole blood in citrate tubes for PCR (may use serum or citrated plasma as a secondary source). • Serum for acute and convalescent antibody titers. • Sputum, if present, for Gram stain, culture, PCR, or direct immunofluorescence stain. If sputum specimen cannot be collected, a throat culture should be obtained. • Pathology specimens for stain, culture, histopathology, and PCR. • Any purulent drainage, if present, for Gram stain, culture, and direct immunofluorescence stain. Collection and Handling of Clinical Specimens Blood Cultures Collect appropriate blood volume and number of sets of blood cultures (usually 2 sets initially and then as clinically indicated). BACTEC adult culture bottles require 10 mL per bottle. If the patient has received antibiotics, blood cultures containing resin to absorb the antibiotics are recommended. Samples should be 0701016F.doc 3-58 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis transported directly to the laboratory at room temperature. They should be held at room temperature until they are placed on the blood culture instrument or incubator. Do not refrigerate. Blood for PCR Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing; transport chilled and not frozen. Collect one 7 mL red top or TT tube for serum for serologies (may refrigerate at 2ºC to 4ºC for 57 days; freeze at -30ºC or lower if > 7 days). Respiratory Specimens Sputum and induced respiratory secretions for culture and PCR should be transported in sterile, screwcapped containers at room temperature. Material to be transported from 2 to 24 hours after collection should be put in a transport container at 2ºC to 8ºC. Swabs (rayon or Dacron only, never cotton) should be moistened with the medium inside the packet, reinserted into the transport package, and transported at room or refrigerator (2ºC to 8ºC) temperature. Nasal Swabs Use rayon or Dacron (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package and moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Skin Lesions Skin lesions and drainage from lesions can be cultured by spreading the tissue or specimen onto a blood agar plate (trypticase soy agar supplemented with 5% sheep blood, BBL [Becton, Dickinson and Company, Franklin Lakes, New Jersey]), and Ashdown’s media, a selective media for B. pseudomallei. While cultures may show colonial morphologic findings of B. pseudomallei, the organism’s identity can be confirmed with biochemical tests (positive oxidase reaction, Simmons citrate) and microscopic morphologic findings [30]. Biopsies Collect biopsy, scraping, swab, or aspirate of ulcer, lymph node, or other involved tissue. For biopsy, place the tissue or scraping in a sterile container. Transport at room temperature for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Treatment of Confimed or Suspected Melioidosis Therapy will vary with the type and severity of the clinical presentation. For a more complete summary of therapy options, refer to review articles by Cheng et al. [15] and White [31]. Although oral therapy with multiple drugs (doxycycline, chloramphenicol, trimethoprim-sulfamethoxazole, and/or quinolones) was used in the past for treating localized disease in nontoxic patients (associated with relapse rates ranging from 10% to 16%), recent studies suggest that relapses may occur less frequently with an initial 2 week course of intensive therapy (involving IV therapy) initiated in conjunction with at least 3 months of oral antibiotic therapy to prevent relapses. Treatment courses of less than 8 weeks have been associated with relapse rates of approximately 25%; treatment courses of 3 months or longer have been associated with lower relapse rates. 0701016F.doc 3-59 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis B. pseudomallei is sensitive to ceftazidime, imipenem, meropenem, amoxicillin-clavulanate acid, piperacillin/tazobactam, doxycycline, trimethoprim-sulfamethoxazole, and chloramphenicol. B. pseudomallei is intrinsically resistant to aminoglycosides and macrolides [33]. Resistance to fluoroquinolones has been observed, with as high as 50% isolates reported to be resistant or with only intermediate sensitivity [33,34]. Quinolones should not be used as primary eradication therapy or in cases in which sensitivity studies show resistance or only intermediate sensitivity. Individuals with melioidosis are recommended to receive intensive therapy with IV antibiotics (minimum of 10–14 days duration) initiated in conjunction with oral eradication therapy (minimum of 12–20 weeks; 6–12 months for deep-seated infections). Ceftazidime, imipenem, meropenem, and amoxicillinclavulanate are bactericidal and are recommended for use in first-line therapy for the initial intensive therapy for melioidosis [32]. A recent study from Australia using an initial course of IV therapy for at least 14 days combined with trimethoprim-sulfamethoxazole was associated with only one failure out of 60 patients infected with trimethoprim-sulfamethoxazole sensitive isolates of B. pseudomallei [13]. While optional, trimethoprim-sulfamethoxazole is generally recommended during intensive therapy, due to the tissue penetration of the drug (particularly in the prostate, central nervous system, and bones). Intensive Therapy IV therapy is recommended initially (intensive therapy) for a minimum period of 10 to 14 days (longer treatments may be required for severely ill patients or deep-seated locations), to be initiated with or without trimethoprim-sulfamethoxazole (oral therapy) [14,35]. Options for initial parenteral treatment include imipenem (25 mg/kg, up to 1 g) every 6 hours, meropenem (25 mg/kg, up to 1 g) every 8 hours, or ceftazidime (50 mg/kg, up to 2 g) every 6 hours combined generally with oral trimethoprimsulfamethoxazole (8/40 mg/kg, up to 320/1600 mg) every 12 hours. Unasyn (3 g every 6 hours in adults) is also an alternative but has a slightly higher rate of treatment failure than ceftazidime. Trimethoprimsulfamethoxazole is recommended to be included in the treatment regimen for neurologic, cutaneous, bone, and prostate infections. Eradication Therapy Oral eradication therapy is generally administered concomitantly with IV therapy. Duration of oral therapy may depend on the extent and severity of disease. Oral therapy for 12–20 weeks may be recommended for disease localized to the lungs, but prolonged therapy (6–12 months) may be required for deep-seated infections (e.g., severe disease and visceral disease, CNS infections, and prostate infections). Trimethoprim-sulfamethoxazole, either alone or in combination with doxycycline is usually the initial recommended treatment of choice for eradication therapy unless the organism is known to be resistant to trimethoprim-sulfamethoxazole [4,5]. Trimethoprim-sulfamethoxazole plus doxycycline as eradication therapy was similar in efficacy as a triple drug regimen (trimethoprim-sulfamethoxazole, doxycycline, and chloramphenicol) [36]. Doxycycline alone is not recommended as first-line eradication therapy as doxycycline monotherapy has been associated with an increased risk of relapse compared to combination drug eradication therapy [11]. An increased risk of relapse has been observed with amoxicillin clavulanate or quinolones when used as eradication therapy compared to trimethoprimsulfamethoxazole, doxycycline, and chloramphenicol. Increased resistance to trimethoprimsulfamethoxazole has been observed in B. pseudomallei strains in Thailand. Drugs for Eradication Therapy First-line drugs for eradication therapy include trimethoprim-sulfamethoxazole (8/40 mg/kg, up to 320/1600 mg) every 12 hours, doxycycline (2.5 mg/kg, up to 100 mg) every 12 hours and/or chloramphenicol. Amoxicillin-clavulanate (500 mg every 8 hours or 875 mg every 12 hours in adults) and quinolones (if organism known not to be resistant or shows only intermediate sensitivity to quinolones) should be used only as second-line therapy because they were found to be less effective in 0701016F.doc 3-60 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis eradication therapy compared to multiple drug oral therapy with trimethoprim-sulfamethoxazole, doxycycline, and choroamphenicol [13]. One study using quinolones for a median of 15 weeks (range 12–40 weeks) for single drug eradication therapy was associated with 29% failure rate (5 failures to respond and 8 relapses) [37]. Sensitivity testing should be performed to aid with therapy. Follow-up Melioidosis responds slowly to treatment, with a median time to defervescence of 9 days. Patients with pulmonary or visceral abscesses may be febrile for several weeks (> 1 month). Sputum cultures may remain positive for a week in spite of treatment with IV antibiotics to which the organism is sensitive [11Currie]. Blood cultures should be negative by the end of the first week of treatment. Enlargement or appearance of an abscess (especially in skeletal muscle) may occur in the first week of treatment and may not necessarily indicate drug resistance. Large abscesses (e.g., prostatic abscesses or skin abscesses), if accessible, normally should be drained [11]. Most other abscesses, including liver and splenic abscesses, will generally resolve with antibiotic therapy. Patients require follow-up at regular intervals for at least 5 years and may require lifelong follow-up. Patients should be alerted to the life-long risk of relapse and the need to alert healthcare providers, particularly when being evaluated for febrile illnesses or other symptomatology that could represent a melioidosis relapse. Environmental Decontamination and Infection Control The organism can be killed with a sporicidal/germicidal agent or 0.5% hypochlorite solution. Health care workers should follow standard precautions when taking care of patients with infections associated with B. pseudomallei. Sexual transmission has been reported in 2 cases and was believed to be related to genitourinary infection (prostatic abscesses) due to B. pseudomallei [14]. Vertical transmission and nosocomial transmission (from contaminated blood-drawing equipment) have been documented [CDC38]. Transmission from an individual with chronic melioidosis to a sibling with diabetes who was serving as a caretaker was also reported [CDC-38]. However, the exact nature of transmission could not be confirmed. Infected individuals should be cautioned concerning the possible transmission from sexual fluids or breast milk [39]. Private-room placement for melioidosis patients is not necessary. References 1. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, and National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed, pp 128–9. Washington DC, U.S. Government Printing Office, 2007. www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5TOC.htm. Accessed December 7, 2007. 2. Centers for Disease Control and Prevention: Imported melioidosis—South /Florida, 2005, MMWR Morb Mortal Wkly Rpt 2006; 55(32):873–76. 3. Sewell DL: Laboratory-associated infections and biosafety, Clin Microbiol Rev 1995; 8(3):389–405. 4. Schlech WF III, Turchik JB, Westlake RE Jr, Klein GC, Band JD, Weaver RE: Laboratory-acquired infection with Pseudomonas pseudomallei (melioidosis), N Engl J Med 1981; 305:1133–5. 5. Green RN, Tuffnell PG: Laboratory-acquired melioidosis, Am J Med 1968; 44:599–605. 6. Ashdown LR: Melioidosis and safety in the clinical laboratory, J Hosp Infect 1992; 21:301–06. 7. Centers for Disease Control and Prevention: Laboratory exposure to Burkholderia pseudomallei— Los Angeles, California, 2004, MMWR Morb Mortal Wkly Rpt 2004; 53(42):988–90. 8. Peacock SJ, Schweizer HP, Dance DA, Smith TL, Gee JE, Wuthiekanun V, DeShazer D, Steinmetz I, Tan P, Currie BJ: Management of accidental laboratory exposure to Burkholderia pseudomallei and B. mallei, Emerg Infect Dis 1998; 14(7):e2. 0701016F.doc 3-61 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis 9. Holland DJ, Wesley A, Drinkovic D, Currie BJ: Cystic fibrosis and Burkholderia pseudomallei: an emerging problem? Clin Infect Dis 2002; 35:e138–40. 10. Cheng AC, Dance DAB, Currie BJ: Letter to the editor (bioterrorism, glanders, and meliodosis), Eurosurveillance 2005; 10(3):pii528. 11. Currie BJ, Fisher DA, Howard DM, Burrow JNC, Lo D, Selva-nayagam S, Anstey NM, Huffam SE, Snelling PL, Marks PJ, Stephens SP, Lum GD, Jacups SP, Krause VL: Endemic melioidosis in tropical Northern Australia: a 10-year prospective study and review of the literature, Clin Infect Dis 2000; 31:981–86. 12. Suputtamongkol Y, Hall AJ, Dance DA, Chaowagul W, Rajchanuvong A, Smith MD, White NJ: The epidemiology of melioidosis in Ubon Ratchatani, Northeast Thailand, Int J Epidemiol 1994; 23(5):1082–90. 13. Gilad J: Burkholderia mallei and Burkholderia pseudomallei: the causative micro-organisms of glanders and melioidosis. Recent Pats Antiinfective Drug Discov 2007; 2(3):233–41. 14. Currie BJ, Fisher DA, Anstey NM, Jacups SP: Melioidosis: acute and chronic disease, relapse and reactivation, Trans R Soc Trop Med Hyg 2000; 94:301–04. 15. Cheng AC, Currie BJ: Melioidosis: epidemiology, pathophysiology, and management, Clin Microbiol Rev 2005; 18(2):383–416. 16. Walsh AL, Smith MD, Wuthiekanun V, Suputtamongkol Y, Desakorn V, Chaowagul W, White NJ: Immunofluorescence microscopy for the rapid diagnosis of melioidosis, J Clin Pathol 1994; 47:377– 79. 17. Dance DA, Wuthiekanun V, Naigowit P, White NJ: Identification of Pseudomonas pseudomallei in clinical practice: use of simple screening tests and API 20NE, J Clin Pathol 1989; 42:645–48. 18. Lowe P, Haswell H, Lewis K: Use of various common isolation media to evaluate the new VITEK 2 colorimetric GN card for identification of Burkholderia pseudomallei, J Clin Microbiol 2006; 44(3):854–56. 19. Wuthiekanun V, Suputtamangkoi Y, Simpson AJH, Kanaphun P, White NJ: Value of throat swab in the diagnosis of melioidosis, J Clin Microbiol 2001; 39(10):3801–02. 20. Tanpiboonsak S, Paemanee A, Bunyarataphan S, Tungpradabkul S: PCR-RFLP based differentiation of Burkholderia mallei and Burkholderia pseudomallei, Mol Cell Probes 2004; 18(2):97–101. 21. Inglis TJ, Aravena-Roman M, Ching S, Croft K, Wuthiekanun V, Mee BJ: Cellular fatty acid profile distinguishes Burkholderia pseudomallei from avirulent Burkholderia thailandensis, J Clin Microbiol 2003; 41(10):4812–14. 22. Wuthiekanun V, Anuntagool N, White NJ, Sirisinha S: Short report: a rapid method for the differentiation of Burkholderia pseudomallei and Burkholderia thalilandensis, Am J Trop Med Hyg 2002; 66(6):759–61. 23. Steinmetz I, Reganzerowski A, Brenneke B, Haussler S, Simpson A, White NJ: Rapid identification of Burkholderia pseudomallei by latex agglutination based on an exopolysaccharide-specific monoclonal antibody, J Clin Microbiol 1999; 37(1):225–28. 24. Inglis TJJ, Merritt A, Chidlow G, Aravena-Roman M, Harnett G: Comparison of diagnostic laboratory methods for identification of Burkholderia pseudomallei, J Clin Microbiol 2005; 43(5):2201–06. 25. Wuthiekanun V, Desakorn V, Wongsuvan G, Amornchai P, Cheng AC, Maharhjan B, Limmathurotsakul D, Chierakul W, White NJ, Day NPJ, Peacock SJ: Rapid immunofluorescence microscopy for diagnosis of melioidosis, Clin Diag Lab Immunol 2005; 12(4):555–56. 0701016F.doc 3-62 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis 26. Miller WR, Pannell L, Ingalls MS: Experimental chemotherapy in glanders and melioidosis, Am J Hyg 1948; 47(2):205–13. 27. Russell P, Eley SM, Ellis J, Green M, Bell DL, Kenny DJ, Titball RW: Comparison of efficacy of ciprofloxacin and doxycycline against experimental melioidosis and glanders, J Antimicrob Chemother 2000; 45:813–18. 28. Batmanov VP, Ilyukhin VI, Lozovaya NA, Andropova NV: Abstract No. 6, Urgent prophylaxis of experimental melioidosis in white rats in Programs and Abstracts Book of the 2004 ASM Biodefense Research Meeting, p 31. Baltimore, MD, American Society of Microbiology, 2004. 29. Sivalingam SP, Sim SH, Jasper LCW, Wang D, Liu Y, Ooi EE: Pre- and post-exposure prophlyaxis of experimental Burkholderia pseudomallei infection with doxycycline, amoxicillin/clavulinic acid and co-trimoxazole, J Antimicrob Chemother 2008; 61:674–78. 30. Wang Y-S, Wong C-H, Kurup A: Cutaneous melioidosis and necrotizing fascitis caused by Burkholderia pseudomallei, Emerg Infect Dis 2003; 9(11):1484–85. 31. White NJ: Melioidosis, Lancet 2003; 361(9370):1715–22. 32. Simpson AJH, Suputtamongkol Y, Smith MD, Angus BJ, Rajanuwond A, Wuthiekanun V, Howe PA, Walsh AL, Chaowagul W, White NJ: Comparison of imipenem and ceftazidime as therapy for severe melioidosis, Clin Infect Dis 1999; 29:381–87. 33. Thibault FM, Hernandez E, Vida DR, Giradet M, Cavallo J-D: Antibiotic susceptibility of 65 isolates of Burkholderia pseudomallei and Burkholderia mallei to 35 antimicrobial agents, J Antimicrob Chemother, 2004; 54:1134–38. 34. Winton MD, Everett ED, Dolan SA: Activities of five new fluoroquinolones against Pseudomonas pseudomallei, Antimicrob Agents Chemother 1988; 32(6):928–29. 35. Webling DD: Genito-urinary infections with Pseudomonas pseudomallei in Australian aboriginals, Trans R Soc Trop Med Hyg 1980; 74(1):138–39. 36. Chaowagul W, Chierakul W, Simpson AJ, Short JM, Stepniewska K, Maharjan B, Rajchanuvong A, Busarawong D, Limmathurotsakul D, Cheng AC, Wuthiekanun V, Newton PN, White NJ, Day NPJ, Peacock SJ: Open-label randomized trial of oral trimethoprim-sulfamethoxazole, doxycycline, and chloramphenicol compared with trimethoprim-sulfamethoxazole and doxycycline for maintenance therapy of melioidosis, Antimicrob Agents Chemother 2005; 49:4020–25. 37. Chaowagul W, Suputtamongkul Y, Smith MD, White NJ: Oral fluoroquinolones for maintenance treatment of melioidosis, Trans R Soc Trop Med Hyg 1997; 91(5):599–601. 38. Centers for Disease Control and Prevention. Meliodosis: General information. http://www.cdc.gov/nczved/divisions/dfbmd/diseases/melioidosis. Accessed October 27, 2010. 39. McCormick JB, Sexton DJ, McMurry JG, Carey E, Hayes P, Feldman RA: Human-to-human transmission of P. pseudomallei, Ann Intern Med 1975; 83(4):512–13. 0701016F.doc 3-63 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis Table 3-12. Diagnostic Testing for Melioidosis Specimen Site and Test Aerobic blood cultures Method of Collection Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Transport Transport at room temperature. No refrigeration. Comments Further sets of blood cultures obtained as clinically indicated. Blood cultures in septicemia are often positive, with organism detected within 48 hours in 90% blood cultures. Automated blood culture methods are typically more rapid. Serum for PCR Serum for acute and convalescent serologies Whole blood is preferable (collected in citrate tube) for PCR; serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for PCR. Transport chilled (not frozen). Collect 1 mL minimum (7 mL adequate) of serum for serology in either red top or TT tubes and spin. Refrigerate at 2ºC to 4ºC for 5 to 7 days. PCR tests, though not validated, may be useful in differentiating among B. mallei, B. pseudomallei, and B. thailandensis. Bacterial fatty acid methyl ester (FAME) profile analysis by gas-liquid chromatography (GLC) to detect a cellular fatty acid profile may be used to distinguish B. pseudomallei from B. thailandensis. Freeze at -30ºC or lower if greater than 7 days. Agglutination tests: Not positive for at least 7 to 10 days; high background titer in normal sera (1:320 to 1:640) makes interpretation difficult. Reproducible fourfold increase in serology is considered diagnostic of infection. A single agglutination titer > 1:160 with compatible clinical picture suggests active infection of melioidosis or glanders. Agglutination titers ≥ 1:40 with a compatible clinical presentation or known negative baseline titer may be significant in persons from nonendemic areas. CF tests more specific but less sensitive and may require 40 days for seroconversion. Often considered positive if titer ≥ 1:20. Follow-up titers recommended for 4 to 6 weeks as delayed seroconversion may occur. Latex agglutination tests (based on poly- or monoclonal antibodies to lipopolysaccharide or exopolysaccharide) reported to be more specific for diagnosis. 0701016F.doc 3-64 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis Table 3-12. Diagnostic Testing for Melioidosis (cont.) Specimen Site and Test Method of Collection Collect > 1 mL of a lower respiratory specimen into a sterile container for stains and culture. Transport Comments Transport in sterile, screw-capped container at room temperature, when transport time is < 2 hours. For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. Sputum culture should be obtained if pneumonia. If no sputum production, throat culture with swab may be positive). Gram stain. Methylene blue or Wright stain may reveal bipolar “safety pin” appearance. Morphology variable in clinical specimens. Will grow in standard media. Selective media (modified Ashdown’s medium) has been used for isolation of organism from nonsterile sites such as sputum and pharyngeal cultures. Addition of 1% to 5% glucose, 5% glycerol, or meat infusion nutrient agar may accelerate growth. Isolation requires 48 to 72 hours in agar at 37.5ºC. Generally grows faster and less fastidious than B. mallei. Sputum for Gram stain, culture, and PCR Direct immunofluorescent microscopy (including rapid tests) of clinical samples (pus, sputum, urine) with 66% sensitivity/ 99% specificity compared to culture. Nasal/throat swab 0701016F.doc Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport packet at room temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. If no sputum production, throat culture maybe positive. 3-65 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Melioidosis Skin lesions and biopsies: PCR, histopathology cultures and stains Skin lesions and drainage from lesions can be cultured by spreading tissue or specimen onto blood agar plate (trypticase soy agar supplemented with 5% sheep blood, BBL [Becton, Dickinson and Company, Franklin Lakes, NJ]) and Ashdown’s media. Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Aspirates: sample should be capped, secured by tape, and sent to lab at room temperature (22°C to 28°C) for immediate processing. Refrigerate (2ºC to 8ºC) if processing will be delayed. Biopsies: Place tissues in sterile containers with 1 to 2 drops sterile nonbacteriostatic normal saline to keep moist. Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. Cultures may show colonial morphologic findings; can be confirmed by biochemical tests (positive oxidase reaction, Simmons citrate), and microscopic morphologic findings. Skin biopsies. 0701016F.doc 3-66 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis BRUCELLOSIS—Br ucella spp Overview of Laboratory Exposures Background Brucellosis is a zoonotic disease caused by an infection with one of six species of Brucella, a group of intracellular, gram-negative coccobacilli. The natural reservoirs for these organisms are sheep, cattle, and goats. In animals, infection with the organisms primarily involves the reproductive tract, causing septic abortions and sterility. Infection is transmitted to humans by direct contact with infected animals or their carcasses, or via ingestion of unpasteurized milk or milk products. Infections in abattoir and laboratory workers suggest that Brucella are highly infectious via the aerosol route, with an estimate of only 10 to 100 bacteria being required to cause clinical disease. Experience with Laboratory Exposures Laboratory-acquired infections with Brucella have been reported to occur by inhalation and percutaneous or conjunctival inoculation. During the U.S. BW program (1943–1969) at USAMRIID (then called the U.S. Army Medical Unit, Fort Detrick), 95 cases of brucellosis were diagnosed. The suspected route of exposure in nearly all cases was aerosol, as most individuals had no history of a specific laboratory accident before the onset of their symptoms. A total of 51 cases were diagnosed during the 6 year period between 1943 and 1949, before the commercial availability of BSCs, which were installed at Fort Detrick beginning in 1950 [1]. While there was a marked decline in cases of brucellosis beginning in 1952, with only 1 case diagnosed in the 11 year period from 1958 to 1969, the decrease may be attributed to a decline in the number of persons working with the agent in addition to the use of BSCs [2]. Infections diagnosed in hospital laboratory workers in brucellosis-endemic areas have been associated mainly with aerosol exposures, resulting from unsafe laboratory practices such as sniffing culture plates and working on open bench tops with viable cultures before identifying the organism as Brucella. However, disease has also been acquired from ingestion of the organism (mostly in older literature, particularly associated with the practice of mouth pipetting) and by self-inoculation. The laboratory attack rates of 30%–100% have been reported and depend on multiple factors, such as the degree of aerosolization and the exposure dose. Laboratory attack rates with ranges from 30% to 100% have been reported [3]. In a single laboratory event (non-USAMRIID laboratory), the accidental breakage of a centrifuge tube containing live B. abortus serotype 1 atypical strain resulted in 12 persons contracting brucellosis 6 weeks to 5 months after exposure. Diagnosis was based on results from serological testing that was performed weekly for 6 months following the event [4]. Nine asymptomatic individuals had seroconversion of titers before the onset of symptoms and were immediately given doxycycline (200 mg daily) and rifampicin (600 mg daily). They experienced symptoms of fever, headache, and chills of only a few days duration. This was in contrast to the 3 symptomatic individuals treated with doxycycline and rifampicin who had a prolonged illness of fever, headache, and chills lasting 2–3 weeks, in addition to symptoms of anorexia, malaise, myalgia, or arthralgia, which lasted an additional 2–3 weeks. Follow-up of the 12 individuals demonstrated maximum antibody titers between 2–3 months post-exposure, with 11 of the 12 individuals having positive titers until nearly a year after the exposure. No relapses were noted over the 8 year followup period, possibly due to the low virulence of this particular strain in humans or, as in the case of the 9 initially asymptomatic individuals, due to the early initiation of antibiotics immediately after seroconversion [4]. 0701016F.doc 3-67 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis In another hospital laboratory incident, 6 laboratory workers were identified as having had a high-risk exposure to B. melitensis because they had sniffed and manipulated cultures outside a BSC while vortexing tubes and preparing subcultures and suspensions for identification and susceptibility testing. Five individuals were given post-exposure prophylaxis for 3 weeks: 4 were given doxycycline 100 mg 2 times daily plus rifampin 600 mg once daily, and 1 pregnant laboratory worker was given trimethoprimsulfamethoxazole 160 mg/800 mg 2 times daily. One individual declined prophylaxis but became symptomatic, and brucellosis was confirmed 10 weeks later. The 5 individuals who received postexposure prophylaxis remained healthy and did not seroconvert. In addition, 20 laboratory personnel who were identified as being at low risk, i.e., worked in the laboratory but did not manipulate or work directly with the organism (with the exception of 1 individual who did not manipulate the culture but did sniff the culture plate on one occasion), remained healthy and did not seroconvert [5]. In 2007, a laboratory preparedness survey (as a training exercise for laboratories for handling specimens that may contain virulent zoonotic agents) resulted in potential exposure to over 900 laboratory workers at over 250 participating laboratories due to failure of the laboratory workers to perform all manipulations of the specimen within a Class II biosafety cabinet (specimen contained RB51 B. abortus [a vaccine strain] but was mislabed as a routine patient specimen). All individuals (74% were assessed as high risk of exposure) were given 21 days post-exposure prophylaxis with doxycycline and rifampin, and no individuals developed brucellosis [6]. Because of the low-infective dose of Brucella, brucellosis continues to be a commonly reported laboratory-associated bacterial infection [7]. Disease severity and clinical presentation vary with the infecting strain of Brucella. B. melitensis is the most virulent species. B. abortus and B. canis are often associated with an insidious onset of disease and frequent relapses and B. suis infection with localized abscess formation. All forms have caused illness in laboratory personnel. Specific Laboratory Hazards and Biosafety Level Working Conditions Cases of brucellosis have occurred commonly in both research and clinical laboratories. Brucella may be present in blood, CSF, semen, and occasionally urine of infected animals or persons. Infections acquired in the clinical laboratory are mainly from inhalation of the organism resulting from the unsafe practice of sniffing culture plates or from manipulating viable organisms on an open bench before the identification of the organism as Brucella [8]. Aerosols generated during laboratory procedures have resulted in large outbreaks of brucellosis [9]. Direct skin contact (perhaps through microabrasions of the skin) with cultures or with infectious clinical specimens from animals (e.g., blood and uterine discharges) is commonly implicated. Mouth pipetting, accidental parenteral inoculations, and mucosal exposures have also resulted in infection [8]. BSL-2 facilities are recommended for human and animal clinical specimens. BSL-3 conditions and ABSL-3 facilities are recommended for all manipulations of cultures of pathogenic Brucella spp and for animal studies [10]. At USAMRIID, all work with Brucella spp is conducted in BSL-3/ABSL-3 containment. Experimental aerosol studies with Brucella spp are conducted in a Class III BSC. 0701016F.doc 3-68 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis The Disease Clinical Features Brucellosis, also known as “undulant fever,” typically presents as a nonspecific febrile illness resembling influenza. After a prodromal period (generally 1 to 7 days), individuals develop an acute or subacute febrile illness, associated with moderate to severe fatigue, myalgia and/or arthralgia, headache, and leukopenia with a relative lymphocytosis [8]. The primary clinical features of brucellosis are listed in Table 3-13. The incubation period is highly variable and difficult to ascertain: usually 5 to 60 days but occasionally several months. Brucellosis may last for several days, months, or occasionally a year or more if not adequately treated. Table 3-13. Clinical Features of Brucellosis Criterion Route of Exposure Incubation Period Clinical Presentation Prognosis Clinical Features Percutaneous or conjunctival inoculation. Ingestion (usually unpasteurized milk or milk products). Inhalation. Highly variable. Generally 7 to 21 days postexposure (usually within 8 weeks); occasionally several months Fever, headache, myalgias, arthralgia, back pain, sweats, chills, anorexia, weight loss, and generalized malaise are common complaints; sometimes misdiagnosed as “flu.” Undulant fever pattern may occur if not treated. Gastrointestinal symptoms (anorexia, nausea, vomiting, diarrhea, and constipation) occur in up to 70% of adult cases. Ileitis, colitis, and granulomatous or mononuclear infiltrative hepatitis may occur, with hepatomegaly and splenomegaly present in 45% to 63% of cases and normal or mildly elevated liver function tests. Osteoarticular complications include bursitis, tenosynovitis, arthritis, osteomyelitis (particularly vertebral osteomyelitis), sacroiliitis, discitis, and paravertebral abscess (20% to 60%). Hematologic symptoms include anemia, leukopenia, or thrombocytopenia. Epididymitis in 20% of males. Neuropsychiatric symptoms include depression, headache, and irritability often in excess of symptom severity. Neurologic symptoms in < 5% of cases, present as acute or chronic meningitis. If untreated, may develop undulant fever pattern. Mortality < 2%, with majority due to endocarditis or meningitis. Acute brucellosis during first and second trimesters of pregnancy has led to spontaneous abortion in up to 4% of women if untreated, and untreated disease may be associated with intrauterine fetal death (2%) with onset of third trimester. Diagnosis A history that elicits potential exposure to Brucella (e.g., laboratories, animals, animal products, or environmental exposure to locations inhabited by potentially infected animals) is an important diagnostic factor to consider in cases of suspected brucellosis. A presumptive diagnosis of brucellosis may be made by serological tests. Although a number of serologic techniques have been developed and tested, including serum agglutination tests for IgM and IgG, the tube agglutination test remains the standard method. This test, which measures the ability of serum to agglutinate killed organisms, reflects the presence of anti-O-polysaccharide antibody. Use of the tube agglutination test (after treatment of serum with 2-mercaptoethanol or dithiothreitol to dissociate IgM into monomers) detects IgG antibody. Most cases of active brucellosis have a titer of ≥ 1:160, and a titer ≥ 1:160 with clinical symptoms of brucellosis is generally considered a presumptive diagnosis for brucellosis. 0701016F.doc 3-69 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis As many patients already have high titers at the time of clinical presentation, a fourfold rise in titer may not occur. IgM rises early in disease and may persist at low levels (e.g., 1:20) for months or years after successful treatment. Persistence or increase of 2-mercaptoethanol-resistant titers has been associated with persistent disease or relapse. Serum testing should always include dilution to at least 1:320 since inhibition of agglutination may occur at lower dilutions. The tube agglutination test does not detect antibodies to B. canis because this rough organism does not have O-polysaccharide on its surface. Although immunoenzymatic assays (e.g., ELISAs) have been developed for use with B. canis and other Brucella spp, they have yet to be validated. Confirmation of a diagnosis of brucellosis can be made by culture (i.e., blood, bone marrow, or other tissue). Cultures should be held for 14 days (28 days in rare circumstances), with weekly subcultures onto solid medium. Because it is extremely infectious for laboratory workers, organisms should be subcultured only in a biohazard hood. The reported frequency of isolation from blood varies widely from < 10% to 90%. B. melitensis is said to be more readily cultured than B. abortus. Culture of bone marrow may increase the yield [11]. PCR assays may be available but have not been validated. Preliminary results have demonstrated a high sensitivity and specificity in the diagnosis of brucellosis, including acute, chronic, and relapsed brucellosis [12]. Management of Potential Exposures to Brucella (Asymptomatic Persons) Initial History and Risk Assessment Individuals with potential exposures to Brucella spp should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure are generally followed (outlined in Chapter 2). Diagnostic Testing Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater-risk exposure generally require the following: – Two 10.0 mL SSTs for research serology and storage at time of exposure (should be within 7 days of exposure). – Two 10.0 mL SSTs for convalescent research serology at 14 days or later post-exposure, if indicated, for serological diagnosis. Must place in isolation bag labeled: “Suspicion of Brucellosis.” – May consider (on a case-by-case basis) nasal swabs for culture for epidemiological purpose if within 24 hours of exposure. A positive nasal swab within 24 hours of exposure only indicates exposure to Brucella, and a negative swab does not exclude exposure. Post-exposure Prophylaxis Males and Non-pregnant Females Post-exposure prophylaxis is indicated for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. Based on limited clinical data (as discussed earlier in this chapter) and experience with using two drugs for the treatment of brucellosis, the current recommendations for post-exposure chemoprophylaxis include a two drug regimen of doxycycline, 100 mg, orally every 12 hours and rifampin, 600 mg (or 450 mg, depending on body 0701016F.doc 3-70 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis weight), orally every 24 hours generally for 3 weeks. Adverse events of nausea, vomiting, and anorexia were commonly reported in one group of 6 individuals receiving post-exposure prophylaxis, with fever and mild hepatitis in one person [13]. While 7-day courses of doxycycline have been used as postexposure prophylaxis in small numbers of persons, there are no data from controlled studies to support the efficacy of a 7-day course with a single drug as prophylaxis [14]. Pregant Females Recommendations for antibiotic prophylaxis in pregnant women with significant risk exposure to Brucella: • Consultation with infectious diseases and/or obstetrics experts (or other experienced health care provider) is highly recommended. The risk of post-exposure prophylaxis must be weighed against the exposure and risk of disease from Brucella. A combination of trimethoprimsulfamethoxazole plus rifampin may be considered. Trimethoprim-sulfamethoxazole may be associated with a theoretical risk of neonatal bilirubinemia when given before delivery. • Ciprofloxacin is a Pregnancy Category C drug. Although the TERIS concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Doxycycline is a Category D agent in pregnancy and may cause discoloration of fetus teeth if given during the last half of pregnancy. The use of these drugs for prophylaxis must be weighed against the risk of disease. Management of Suspected Brucellosis (Symptomatic Persons) Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to Brucella must be documented with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Blood for culture (hold cultures for at least 14 days) • Whole blood in citrate tubes for PCR (may use serum or citrated plasma as a secondary source) • Serum for immunoassays (acute and convalescent) • Tissue (i.e., bone marrow) for culture and histopathology Collection and Handling of Clinical Specimens Blood Cultures Collect appropriate blood volume and number of sets of blood cultures (usually 2 sets initially) and then as clinically indicated. BACTEC adult culture bottles require 10 mL per bottle (pediatric 5 mL per bottle). If the patient has received antibiotics, blood cultures containing resin to absorb the antibiotics are recommended. Transport directly to laboratory at room temperature. Hold at room temperature until placed on the blood culture instrument or incubator. Do not refrigerate. Note: Blood cultures should be maintained in a closed system, and clinical isolates from blood or any other site should be handled in BSL-3 cabinets. Follow established laboratory protocol for processing blood cultures. Blood for PCR 0701016F.doc 3-71 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis Collect a minimum of 3 mL of whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing; transport chilled and not frozen. Collect one 7 mL red top or TT tube for serum for serologies (may refrigerate at 2ºC to 4ºC for 5 to 7 days; freeze at -30ºC or lower if > 7 days). Nasal Swabs Use rayon or Dacron (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2Cº to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Biopsies Collect biopsy (i.e., bone marrow or other involved tissue). For biopsy, place the tissue or scraping in sterile container. Transport at room temperature for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Treatment of Confirmed or Suspected Brucellosis Therapy will vary with the type and severity of the clinical presentation. • • • • • Most effective proven treatment for acute brucellosis has been a combination of tetracycline (500 mg orally every 6 hours for 6 weeks) and streptomycin (1 g IM daily for the first 2–3 weeks [15,16]. This regimen has been replaced doxycycline (100 mg orally 2 times daily for 45 days) and gentamicin (5 mg/kg/day IM for 7 days) [17,18]. This regimen was found to be an equally effective (94.8% cure rate) as doxycycline for 45 days and streptomycin for 14 days (92.6% cure rate) in one study [19]. Doxycycline (100 mg orally 2 times daily for 6 weeks) in combination with rifampin (600 to 900 mg orally once daily for 6 weeks has also been effective in treatment of uncomplicated brucellosis (may be associated with a higher relapse rate than doxycycline with an aminoglycoside) [18]. Ofloxacin or ciprofloxacin plus rifampin or trimethoprim-sulfamethoxazole plus rifampin may be alternatives if primary treatment cannot be given, but may be less effective [12,18]. Trimethoprim-sulfamethoxazole and rifampin may be considered for treatment of brucellosis in children and pregnancy. Relapse rates are 5%–10% for most combination oral regimens and higher with monotherapy (up to 30% with trimethoprim-sulfamethoxazole alone). Acute, complicated brucellosis often requires long-term, triple-drug therapy. A combination of oral rifampin and doxycycline plus IM streptomycin for the first 2–3 weeks has been used most frequently. Skeletal disease may require 6–8 weeks of antibiotics. Patients who have meningoencephalitis and endocarditis should receive at least 90 days of therapy and may require more than 6 months of therapy. Endocarditis usually requires valve replacement. Necrotizing orchitis and other suppurative complications may require surgical excision or drainage. Environmental Decontamination and Infection Control Brucella may survive for prolonged periods in the environment (6 weeks in dust and 10 weeks in soil or water). Contact surfaces that are free of organic matter can be decontaminated with a 0.5% hypochlorite solution; higher concentration or other disinfectants for gram-negative microorganisms should be used where organic matter cannot be effectively reduced or controlled. 0701016F.doc 3-72 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis Health care workers should follow standard precautions when taking care of patients with infections associated with Brucella. The agent may be present in blood, CSF, semen, and occasionally urine. Brucellosis is spread readily via bodily fluids and aerosols. Person-to-person transmission has been reported via tissue transplantation and sexual contact. Women who are breast feeding may transmit the infection to their infants. References 1. Wedum AG: The Detrick experience as a guide to the probable efficacy of P4 microbiological containment facilities for studies on microbial recombinant DNA molecules, J Am Biol Safety Assoc 1996;1(1):7–25. 2. Rusnak JM, Kortepeter MG, Hawley RJ, Anderson AO, Boudreau E, Eitzen E: Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers, Biosec Bioterr Biodef Strat Prac Sci 2004;2(4):1–13. 3. Meyer KF, Eddie B: Laboratory infections due to Brucella, J Infect Dis 1941;68:24–32. 4. Fiori PL, Mastrandrea S, Rappelli P, Cappuccinelli P: Brucella abortus infection acquired in microbiology laboratories, J Clin Micro 2000;38:2005–06. 5. Robichaud S, Libman M, Behr M, Rubin E: Prevention of laboratory-acquired brucellosis, Clin Infect Dis 2004;38:e119–22. 6. CDC: Update: potential exposures to attenuated vaccine strain Brucella abortus RB51 during a laboratory proficiency test—United States and Canada, 2007, MMWR Morb Mortal Wkly Rep 2008;57(02):36–39. 7. Della-Porta AJ, Murray PK: Management of biosafety, In Richmond JY, ed: Anthology of Biosafety: Perspectives on Laboratory Design, pp 1–23. Mundelein, IL, American Biological Safety Association, 1999. 8. Howe C, Miller ES, Kelly EH, Bookwalter HL, Wllingson HV: Acute brucellosis among laboratory workers, N Engl J Med 1947; 236:741–47. 9. Skinhölj P. Occupational risks in Danish clinical chemical laboratories: II infections, Scan J Clin Lab Invest 1974; 33(1):27–29. 10. Yagupsky P, Baron EJ: Laboratory exposures to brucellae and implications for bioterrorism, Emerg Infect Dis 2005; 11(8):1180–85.Washington State Department of Health: Surveillance and reporting guidelines for brucellosis. http://www.doh.wa.gov/notify/guidelines/brucellosis.htm. Accessed December 7, 2010. 11. Gotuzzo E, Carrillo C, Guerra J, Llosa L: An evaluation of diagnostic methods for brucellosis – The value of bone marrow culture, J Infect Dis 1986; 153(1):122–25. 12. Nimri LF: Diagnosis of recent and relapsed cases of human brucellosis by PCR assay, BMC Infect Dis 2003; 3:5. 13. Maley MW, Kociuba K, Chan RC: Prevention of laboratory-acquired brucellosis: significant side effects of prophylaxis, Clin Infect Dis 2006; 42(3):433–34. 14. Zervos MJ, Bostic G: Exposure to Brucella in the laboratory, Lancet 1997; 349(9052):651. 15. Acocella G, Bertrand A, Beytout J, Durrande JB, Garcia Rodriguez JA, Kosmidid J, Micoud M, Rey M, Rodriguez Zapata M, Roux J, Stahl J-P: Comparison of three different regimens in the treatment of acute brucellosis: a multicenter multinational study, J Antimicrob Chemother 1989; 23(3):433–39. 16. Ariza J, Gudiol F, Pallares R, Viladrich PF, Rufi G, Corredoira J, Miravitlles MR: Treatment of human brucellosis with doxycycline plus rifampin or doxycycline plus streptomycin: a randomized, double-blind study, Ann Intern Med 1992; 117(1):25–30. 0701016F.doc 3-73 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis 17. Solera J, Geijo P, Largo J, Rodriguez-Zapata M, Gijón J, Martinez-Alfaro E, Navarro E, Macia MA, Grupo de Estudio de Castila-la Mancha de Enfermedades: A randomized, double-blind study to assess the optimal duration of doxycycline treatment for brucellosis, Clin Infect Dis 2004; 39(12):1776–82. 18. Skalsky K, Yahav D, Bishara J, Pitlik S, Leibovici L, Paul M: Treatment of human brucellosis: systematic review and meta-analysis of randomised controlled trials, Br Med J 2008; 336(7646):701– 04. 19. Roushan MRH, Mohraz M, Hajiahmadi M, Ramzani A, Valayati AA: Efficacy of gentamicin plus doxycycline versus streptomycin plus doxycycline in the treatment of brucellosis in humans, Clin Infect Dis 2006; 42:1075–80. . 0701016F.doc 3-74 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Brucellosis Table 3-14. Diagnostic Testing for Brucellosis Specimen Site and Test Aerobic blood cultures Serum for PCR Serum for agglutination and tube agglutination test for acute and convalescent serologies Nasal/throat swab Biopsy specimens (i.e., bone marrow, liver) 0701016F.doc Method of Collection Transport Comments Collect blood cultures (usually 2 sets initially). BACTEC cultures require 10 mL per bottle. Use blood cultures containing resin if patient has received prior antibiotics. Transport at room temperature. No refrigeration. Further sets of blood cultures obtained as clinically indicated. Whole blood is preferable (collected in citrate tube) for PCR; serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for PCR. Transport chilled (not frozen). PCR is investigational. Collect minimum of 1 mL (7 mL adequate) serum for serology in either red top or TT tubes and spun. Refrigerate at 2ºC to 4ºC for 5 to 7 days. Agglutination tests: A titer ≥ 1:160 considered diagnostic. Most patients have high titers at time of clinical presentation so fourfold rise in titer may not occur. Caution: Blood cultures should be maintained in a closed system and clinical isolates from blood or other sites should be handled under BSL-3. Freeze at -30ºC or lower if greater than 7 days. IgM rises early in disease and may persist at low levels (e.g., 1:20) for months or years after successful treatment. Use rayon or Dacron (not cotton) swabs. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport packet at room temperature. Biopsies: Place tissues in sterile containers. Biopsies. Transport at room temperature for immediate processing. Keep the specimen chilled if processing of the specimen will be delayed. Refrigerate at 2ºC to 8ºC if processing is delayed. Caution: Lab should be notified of culture of possible brucellosis. Clinical isolates from blood or other sites should be handled under BSL-3. Caution: Lab should be notified of culture of possible brucellosis. Clinical isolates from blood or other sites should be handled under BSL-3. 3-75 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Q FEVER—Coxiella burnetii Overview of Laboratory Exposures Background Q fever is a zoonotic disease caused by a rickettsia, C. burnetii. A broad range of domestic (sheep, cattle, and goats) and wild mammals are natural hosts for Q fever and may serve as potential sources of infection for laboratory and animal care personnel. Infection most commonly occurs through airborne transmission of C. burnetii from dried excreta of infected animals or indirect contact (e.g., contaminated dust, straw, wool, manure, or clothing with animal excretions) with infected animals. Exposure to infected, often asymptomatic, sheep and their birth products is a documented hazard to personnel. Individuals at greatest risk are those who work in establishments that process infected animals or their byproducts or work in necropsy rooms, farmers, and abattoir workers. Symptomatic disease in humans most commonly presents as a self-limited febrile illness of 2 to 14 days duration. C. burnetii may also cause chronic infections such as endocarditis or granulomatous hepatitis. Individuals with valvular heart disease and pregnancy are at higher-risk for chronic Q fever. Endocarditis has been reported as high as 39% of infected persons with valvular heart disease [1]. Complications of pregnancy (including spontaneous abortion, stillbirth, intrauterine growth retardation, and premature delivery) have been reported in 81% of pregnant women not given long-term cotrimoxazole, and are highest if infected during the first trimester of pregnancy [2]. Pregnant women, particularly those infected within the first six months of pregnancy, are at risk for chronic Q fever infection (11/17 (65%) cases) [3]. High-risk individuals for developing chronic Q fever should be cautioned concerning work with C. burnetii. The infectious dose of virulent Phase I organisms in laboratory animals has been calculated to be as small as a single organism [4], and airborne particles containing organisms may be carried downwind for 1/2 mile or more. Experience with Laboratory Exposures Q fever was a commonly reported laboratory-associated infection, with outbreaks in several institutions involving 15 or more persons [5]. Between 1950 and 1965, during the U.S. BW program (1943–1969) at Fort Detrick, called the U.S. Army Medical Unit, Fort Detrick, 55 cases of Q fever were documented. All but 1 were felt to be acquired by aerosol exposure [6,7]. Even with the installation of BSCs in the early 1950s, infections continued to occur, most likely related to the low-infective dose and to the organism’s ability to survive in the environment for prolonged periods of time [7]. A review of the initial 50 cases of Q fever diagnosed during the U.S. BW program showed a risk factor of a known exposure in only 5 laboratory workers and a risk factor of working with the agent but no known accidental exposure in 16 persons. The majority of cases (29 of 50 cases) occurred in persons who did not work directly with the agent and had no known identifiable exposure incident, including 13 non-laboratory workers such as clerical workers, engineering personnel, carpenters, janitors, repairmen, steam fitters, and animal handlers [6]. In 1952, Q fever was diagnosed in the spouse of a scientist who worked with C. burnetii at Fort Detrick. The agent was probably acquired from exposure to the organism on her husband’s scalp or hair. Although he always showered and changed clothing before exiting the laboratory, he did not always wash his hair (his head was uncovered while in the laboratory) [7]. After introduction of the formalin-inactivated whole cell Q fever vaccine [6] at Fort Detrick in 1965, Q fever infections decreased from an average of 3.4 cases per year to only 1 infection during the subsequent 40 years [7]. The 1 confirmed case of Q fever diagnosed at Fort Detrick after the introduction of the investigational formalin-inactivated Q fever vaccine resulted from an inhalational exposure due to a breach in the BSC filter. This individual, vaccinated 5 months before exposure, worked with high 0701016F.doc 3-76 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever concentrations of C. burnetii. He presented with a nonspecific flu-like illness [8,9]. Diagnosis of Q fever was confirmed by a rise in serological titers, and symptoms resolved with a regimen of doxycycline. Outbreaks of Q fever have been commonly reported at other laboratories, including the National Institute of Health where 153 cases occurred over 54 days (one death) in 1940 and 47 cases in a subsequent outbreak in 1947. Both outbreaks resulted from aerosol exposure. The second outbreak was associated with aerosolization from preparation of yolk sac antigens from infected eggs, with all infected persons having been in the laboratory within the past 28 days (3 persons only in the building for a short time). Individuals who developed Q fever infection during the 1940 outbreak did not develop symptoms in the second outbreak in 1947 [10,11]. The stability of the organism has been reported to even result in transmission of disease to laundry workers who acquired the disease from contaminated laboratory laundry [12]. Outbreaks of Q fever in laboratory workers have also occurred from working with either naturally or experimentally infected animals (particularly pregnant sheep) [13,14]. An outbreak of Q fever at a university research center in California involved 11 confirmed and more than 30 presumptive cases of Q fever (including one death) over a 3 month period in 1979 due to infected pregnant ewes [15]. A subsequent serologic survey of 580 employees revealed 114 subjects (19.6%) with complement fixation (CF) ≥ 1:8 to C. burnetii. The highest prevalence of CF antibodies was in animal technicians and cage cleaners, as well as individuals who worked on the same floor of the laboratory where studies on the infected sheep were performed. Also 5 of the 9 employees who worked with soiled linen at the campus laundry had elevated CF titers. Additionally, many individuals who had no direct contact with the sheep, but were in the hallways or elevators where the sheep were transported in open carts had elevated CF titers. The sheep were subsequently moved to a containment laboratory suite with negative air pressure, air locks, and high-efficiency particulate filters on exhaust ducts. Specific Laboratory Hazards and Biosafety Level Working Conditions Infectious aerosols are the most common source of infection in laboratory and animal care personnel, but infection may also be acquired by intradermal inoculation (e.g., needlesticks) and less commonly from ingestion [7,16,17]. The potential risk for laboratory exposure to C. burnetii is often higher than for most other agents because of the need to use embryonated eggs or cell culture techniques to propagate C. burnetii, requiring extensive purification procedures and consequently longer exposure to the agent. The agent may be present in the blood, urine, feces, milk, and tissues of infected animals or human hosts and in infected arthropods (mainly ticks) [16,18-20]. The placenta of infected sheep may contain as many as 109 organisms per gram of tissue and milk may contain 105 organisms per milliliter [19,21,22]. BSL-2 facilities are recommended when working with C. burnetii for nonpropagative laboratory procedures, including serological examinations and staining of impression smears. The organism should be handled under BSL-3 conditions for activities involving inoculation, incubation, and harvesting of embryonated eggs or tissue cultures, necropsy of infected animals, and manipulation of infected tissues. Infected rodents should be maintained under ABSL-3 conditions [18]. At USAMRIID, work with C. burnetii is conducted in BSL-3 containment with enhancements (recommended vaccination with the IND Q fever vaccine, mandatory clothing change, use of personal protective equipment, mandatory exit shower, HEPA filtration of exhaust air, and sewage sterilization). Animals experimentally infected with C. burnetii are maintained in ABSL-3 facilities with enhancements. All experimental aerosol studies of C. burnetii are conducted in a Class III BSC. 0701016F.doc 3-77 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever A Q fever vaccine is licensed in Australia but not in the United States. A protocol exists for administration of an investigational Phase 1 formalin-inactivated, freeze-dried Q fever vaccine derived from pathogen-free chicken eggs to at-risk personnel at USAMRIID. Vaccination with a single 0.5 cc SC dose of this killed suspension of C. burnetii provides protection against naturally occurring Q fever and greater than 95% protection against aerosol exposure. Potential protection may be lifelong. Note: Pre-existing immunity to Q fever and increased risk for severe reactions to Q fever vaccination can be assessed with an intradermal (0.1 cc) skin test (Figure 3-5) of Q fever vaccine antigen, which is a 1:300 dilution of the vaccine. Individuals are cleared for laboratory entry 3 weeks after vaccination. Because of the risk of endocarditis from C. burnetii, the CDC has recommended that individuals with valvular heart disease not work with C. burnetii [23]. Figur e 3-5. Positive Q Fever Skin Test The Disease Clinical Features Q fever generally presents as an acute, undifferentiated fever 1 to 3 weeks after exposure to C. burnetii (Table 3-15). The incubation period varies according to the number of organisms inhaled. Infection with greater numbers of organisms will result in shorter incubation periods. Q fever is often a self-limited, febrile illness (lasting 2 to 14 days duration) but can also commonly present as pneumonia. Initial symptoms are a sudden onset of high fever, chills, myalgia, severe headache, malaise, weakness, and anorexia. Between 4 and 6 days after the onset of fever, radiographic and ausculatory signs of pneumonia may be seen in approximately 50% of patients. Q fever will vary in severity. For example, of the initial 50 laboratory-acquired infections from C. burnetii diagnosed during the U.S. BW program, 30 individuals required hospitalization and 10 were managed as outpatients [6]. In these individuals, a Q fever diagnosis was made by retrospective serological testing, with 4 patients unable to recall symptoms of any recent infectious illness. Of the 40 cases actively evaluated, onset of illness was sudden in 31 and insidious in 9. Nonspecific findings of fever, chills, headache, myalgia, and malaise were present in the majority of these 40 cases. A total of 23 individuals reported having a cough, and 18 complained of chest pain, with the pain described as pleuritic in 10 persons. Pneumonitis was observed in 28 patients, with infiltrates described mainly as oval, wedge shaped, and/or perihilar. However, chest findings on physical exam were minimal except for the occasional presence of fine rales. Infiltrates from Q fever are most commonly nonsegmental (multiple rounded or oval shaped opacities may be observed in many cases) or segmental pleural-based opacities. Pleural effusions are observed in approximately one-third of cases; hilar adenopathy may also be present. 0701016F.doc 3-78 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Table 3-15. Clinical Features of Q Fever Criterion Route of Exposure Incubation Period Clinical Presentation Acute Inhalation of C. burnetii. Chronic Inhalation of C. burnetii. Inoculation (e.g., needle sticks). Possibly inoculation. 7 to 21 days (mean 15, range 2 to 41). 1 to 20 years. Acute nondifferentiated febrile illness, with sudden onset high fever, chills, myalgia, headache (generally severe), malaise, weakness, anorexia. Can present as chronic hepatitis, culturenegative endocarditis, aseptic meningitis, encephalitis, and osteomyelitis. 50% with nonproductive cough. Endocarditis (< 1% after acute infection), generally involving aortic heart valve. No rash. 25% pleuritic chest pain. 20% have may develop chronic fatigue syndrome after infection [23]. Infection can last up to 6 months. Asymptomatic infection felt to occur. Prognosis Rarely fatal CFR ≤ 65%. CFR = Case fatality rate Chronic Q fever, characterized by infection that persists for more than 6 months, is an uncommon but much more serious disease [24]. Patients who have had acute Q fever may develop the chronic form as long as 20 years after initial infection. Endocarditis is the most common form of chronic infection with Q fever (occurring in approximately 1% of persons following Q fever [1]), but chronic infection may also involve infected aneurysms or vascular grafts, hepatitis, osteomyelitis, interstitial pulmonary fibrosis, or meningoencephalitis [16]. Individuals with valvular heart disease are at risk for Q fever endocarditis (reported as high as 39%) [1] and should be cautioned concerning work with C. burnetii. Tansplant recipients, patients with cancer, and those with chronic kidney disease are also at risk of developing chronic Q fever. Pregnant women, particularly those infected within the first three months of pregnancy, are at risk for chronic infection (reported as high as 80% of cases) [2,3]. Complications of pregnancy are highest if infected during the first trimester of pregnancy, and have been reported in 81% of pregnant women not given long-term cotrimoxazole [2]. Q fever may result in spontaneous abortions, growth retardation, premature deliveries, and stillbirths [2,3]. In a recent report on 17 pregnant women with Q fever (11 women developed Q fever during the first trimester and 3 each in the second and third trimesters), fetal death occurred in 8 cases and premature delivery or low birth weight in 9 cases [3]. C. burnetii was detected in the placenta by culture and/or PCR in 4 women who did not receive long-term antibiotics and in 2 of the 4 women who received long-term trimethoprim-sulfamethoxazole. Abortion occurred in only one of the 6 women infected during the second or third trimesters of pregnancy. Approximately 20% of Q fever cases may subsequently develop symptoms of chronic fatigue syndrome [23]. Differential diagnosis of Q fever includes tularemia and pneumonia caused by Mycoplasma pneumoniae, Legionella pneumophilia, Chlamydia psittaci, and C. pneumoniae. 0701016F.doc 3-79 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Diagnosis Diagnosis is most commonly accomplished by serologic testing for antibody titers (high titers to Phase II antigen in acute infection and high titers to Phase I antigen in chronic infection). A number of serologic methods, including indirect immunofluorescent assay (IFA), complement fixation (CF), microagglutination, and ELISA, have been used to diagnose C. burnetii infection [25]. Significant antibody titers are usually not identifiable until 2 to 4 weeks into the illness (approximately 90% patients have detectable antibodies by the third week) [25]. A fourfold rise in titers is considered diagnostic for Q fever by IFA, CF, ELISA, or microagglutination [25-27]. Anti-phase II IgM titer ≥ 1:50 or anti-phase II IgG titer ≥ 1:200 by IFA are predictive of acute Q fever infection [28,29]. IgM titers generally persist for 6 to 8 months [29], but have been reported to persist for a year or longer in 3% of persons with Q fever. An anti-phase I IgG titer of ≥ 1:800 has been found to be predictive and sensitive for diagnosis of chronic Q fever in Europe [1,25,28-30]. However, this cutoff titer should be determined for each geographical area and may not be be applicable for Q fever assays performed in the United States [25]. In diagnosis of chronic Q fever, serial serologic tests should demonstrate an increase in anti-phase I IgG over time in conjunction with a decline or constant levels of anti-phase II IgG titers and evidence of inflammatory disease [CDC web page]. Cross-reactivity of Q fever serological tests has been reported with Bartonella henselae and quintana and Legionella pneumophila [31]. Complement fixation (CF) titers, while specific, may lack sensitivity for diagnosis of Q fever and may be associated with false negative results in chronic Q fever due to a prozone phenomenon effect and with false positive results due to cross-reaction with hen egg antigens [25,29, 32, 33]. A CF titer ≥ 1:32 or ≥ 1:40 to phase II antigen is considered to be predictive for Q fever infection and a CF titer ≥ 1:200 to phase I antigen for chronic Q fever [25, 34]. For ELISA testing (in addition to a fourfold rise in titer), antiphase II IgG ≥ 1,024, anti-phase II IgM ≥ 512, and anti-phase I IgG ≥ 128 have been proposed as cutoff values for Q fever infection [35]. Persistence of anti-phase II IgM by ELISA tests was noted in individuals, even as long as 5 years later in 2 of 8 individuals. Diagnosis of Q fever is generally not accomplished by culture, as culture of C. burnetii is potentially hazardous to laboratory personnel and requires animal inoculation or cell culture. PCR test (blood or tissue) and DFA are available at the CDC for diagnosis of acute infection. For potential aerosolized laboratory exposures, nasal swabs may be obtained for PCR and culture to assess for exposure. Immunofluorescent stain or electron microscopy of biopsy specimens also can demonstrate the organism. Management of Potential Exposure to C. burnetii (Asymptomatic Persons) Documenting the Exposure Initial History and Risk Assessment Individuals with potential exposures to C. burnetii should have the risk of exposure and disease assessed after obtaining a detailed history of the laboratory incident and clinical evaluation of the individual as outlined in Chapter 2. While the decision for post-exposure prophylaxis is determined on a case-by-case basis, guidelines for recommending post-exposure prophylaxis based on risk of exposure are generally followed (outlined in Chapter 2). Diagnostic Testing • Individuals with negligible-risk exposure generally only require observation. • Individuals with minimal- or greater-risk exposure generally require the following: 0701016F.doc 3-80 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever – – – Two 10.0 mL SSTs for research serology and storage at time of exposure (should be within 7 days of exposure). Two 10.0 mL SSTs for convalescent research serology at ≥ 14 days post exposure, if indicated, for serological diagnosis. May consider (on case-by-case basis) nasal swabs for epidemiological purposes if within 24 hours of exposure. A positive nasal swab only indicates exposure to C. burnetii, and a negative swab does not exclude exposure. Post-exposure Prophylaxis Males and Non-pregnant Females Post-exposure prophylaxis is indicated for persons who may have been exposed to the agent but do not have symptoms or evidence of infection on medical evaluation. Post-exposure prophylaxis with doxycycline, 100 mg, orally every 12 hours, or tetracycline, 500 mg, orally every 6 hours, should be initiated following moderate- to high-risk exposure to C. burnetii. Chemoprophylaxis begun too early in the incubation period may delay but not prevent the onset of symptoms. (If prophylaxis is started 1 day post exposure and continued for 5 days, clinical disease has been shown to occur approximately 3 weeks after discontinuation of therapy. If prophylaxis is started 8 to 12 days after the exposure and continued for 5 to 7 days, clinical disease has not been observed after discontinuation of antibiotics [36].) The current prophylaxis recommendation for Q fever is a 7-day course of antibiotics, starting between Day 8 and Day 12 after exposure. There is no data concerning the efficacy of longer duration of post-exposure prophylaxis (i.e., 14 days) when initiated immediately after exposure. There are no data concerning the effectiveness of postexposure prophylaxis for Q fever other than doxycycline. In vitro sensitivity testing has shown the organism to be most sensitive to rifampin and quinolones (difloxacin, oxolinic acid, and ciprofloxacin), and also (although somewhat less effective) to doxycycline, trimethoprim, and chloramphenicol [37]. While treatment failures have been reported with erythromycin and erythromycin was noted to be less effective than doxycycline in the treatment of Q fever, macrolides have resulted in earlier resolution of fever than β-lactam antibiotics [38]. Pregnant Females Recommendations for antibiotic prophylaxis in pregnant women with significant risk exposure to C. burnetii: • • Consultation with infectious diseases and/or obstetrics specialists (or other experienced health care provider) is highly recommended. The risk of post-exposure prophylaxis must be weighed against the exposure and risk of disease from C. burnetii. Complications from Q fever have been reported in as high as 81% of women infected during the first trimester of pregnancy [2]. Risk of Q fever during pregnancy include spontaneous abortion (26%), intrauterine fetal death (5%), premature delivery (45%), or intrauterine growth retardation (5%) [2]. Transplacental infection of the fetus in utero may occur [2,3]. Chronic Q fever infection was reported to occur in as high as 80% of women infected during the initial three months of pregnancy [2,3]. Long-term trimethorpim-sulfamethoxazole in pregnant women with Q fever has been associated with a decrease in the risk of obstetric complications (see treatment of Q fever). While rifampin and trimethoprim-sulfamethoxazole may be used in pregnancy, they do not have bactericidal effects on C. burnetii and there is no data concerning use of these agents for postexposure prophylaxis. Trimethoprim-sulfamethoxazole may be associated with a theoretical risk of neonatal bilirubinemia when given before delivery. Doxycycline is a Category D agent in pregnancy (may cause discoloration of fetus teeth if given during the last half of pregnancy). 0701016F.doc 3-81 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Ciprofloxacin is a Pregnancy Category C drug. Although the TERIS concluded from a review of published data on experiences with ciprofloxacin use during pregnancy that therapeutic doses during pregnancy are unlikely to pose a substantial teratogenic risk, data are insufficient to state that there is no risk. Management of Suspected Q Fever Disease Documenting the Exposure Risk, Clinical History, and Physical Exam The guidelines for documenting a laboratory exposure at USAMRIID are presented in Chapter 2 of this manual. If not previously documented, any potential exposure to C. burnetii must be documented with the information listed in these guidelines. A clinical history and physical exam should be performed. Diagnostic Testing • Serum for acute and convalescent research serology (significant antibody levels may not be identifiable until week 2 or 3 of illness). • Whole blood in citrate tubes for PCR (may use serum or citrated plasma as a secondary source). (Samples should be taken in the acute-phase and prior to antibiotic therapy.) • Impression smears from infected humans or animals can be stained with Giemsa or Gimenez stains or processed for direct immunofluorescent detection of C. burnetii. • Clinical samples of affected tissues can be submitted as aspirates or tissue biopsies for use in IHC, PCR, or culture procedures.. Collection and Handling of Clinical Specimens Blood for Serology and PCR Collect a minimum of 3 mL whole blood (7 mL adequate) in citrate tubes (may use serum or citrated plasma as alternate secondary source) for PCR and antigen testing; transport chilled and not frozen. Collect one 7 mL red top or TT tube for serum for serologies (may refrigerate at 2ºC to 4ºC for 5 to 7 days; freeze at -30ºC or lower if > 7 days). Avoid multiple freeze–thaw cycles by freezing aliquots of the sample. Samples should be stored in aliquots of at least 100 µL in O-ring screw-cap vials to avoid evaporation. The samples may be preserved by adding thimerosal to a final concentration of 0.01%. Nasal Swabs Use rayon or Dacron (never cotton) swabs for nasal or lesion specimens. If using a swab transport carrier, reinsert the swab into the transport package, moisten the swab fabric with the transport medium inside the packet. Transport at room or refrigerator (2ºC to 8ºC) temperature. Refrigerate at 2ºC to 8ºC if processing is delayed. Biopsies Collect biopsy. For biopsy, place the tissue or scraping in sterile container. Transport at room temperature for immediate processing. Refrigerate at 2ºC to 8ºC if processing is delayed. Cultures Cultures are generally not performed for diagnosis of Q fever. Collect 10 mL of blood in sterile syringe and transport immediately to the laboratory (syringe should be capped) at room temperature (refrigerate if > 1 hour for transport). The laboratory will culture by either animal inoculation or cell culture. Cultures should be performed at BSL-3 conditions. 0701016F.doc 3-82 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Specimens for DFA Specimens for DFA should be transported in sterile, screw-capped containers or three sets of slides should be prepared before shipping. Smears should be made promptly when possible. A smear should not be too thick; newsprint should be visible through the dried smear. Slides should be dried well and fixed. Samples that are left at room temperature may develop heavy bacterial or fungal contamination and may render the sample unsuitable for testing. Treatment of Confirmed or Suspected Q Fever Treatment will vary with the type (acute versus chronic Q fever) and severity of the clinical presentation. Most acute cases of Q fever eventually resolve without antibiotic treatment, but all suspected cases should be treated to reduce risks of complications. Doxycycline is recommended as the treatment of choice for acute Q fever (usually 2 to 3 weeks duration), but quinolones or macrolides may be considered as an alternative treatment if doxycline is contraindicated [27,39]. However, studies in recent years have demonstrated failures with doxycycline (given for 2 weeks to 6 months) in the prevention of chronic Q fever in individuals with valvular heart disease [1]. In vitro testing demonstrated the greatest sensitivity to quinolones and rifampin, with chloramphenicol, doxycycline, and trimethoprim also effective [37]. A placebo-controlled, double-blinded study demonstrated earlier resolution of fever with doxycycline compared to erythromycin (3 ± 1.6 days versus 4.3 ± 2 days) [20]. Other retrospective reviews of Q fever patients in humans suggest that doxycycline and erythromycin resulted in earlier resolution of fever (mean of 3 days and 4.26 days, respectively) compared to β-lactam antibiotics (mean of 6.8 days) [40]. In another retrospective analysis of treatment of Q fever, resolution of fever occurred somewhat earlier with doxycycline and azithromycin (mean duration of fever of 2 and 2.5 days, respectively) versus 3.5 days with other antibiotics [41]. Individuals diagnosed with Q fever who have valvular heart disease are at risk for endocarditis (as high as 39% in one study) [1]. In one case series, endocarditis occurred more frequently in infected individuals with valvular heart disease who did not receive antibiotics (6 of 8 persons) or received single-drug treatment with doxycycline for 2 weeks to 6 months (5 of 10 persons), but did not occur in 12 individuals who received both doxycycline and hydrochloroquine for 1 to 15 months [1]. Based on this information, it was recommended that individuals with valvular heart disease diagnosed with Q fever infection be treated with hydroxychloroquine and doxycycline for a year with close follow-up for development of chronic Q fever (including echocardiograms) [1]. However, because mild mitral regurgitation may be seen in as many as 10% of healthy, young adults (who are otherwise thought to be low risk for endocarditis) and chronic Q fever was not observed in the initial Q fever cases in U.S. military troops in 2003 and 2004, clinical and serological follow-up has been recommended for a minimum of 2 years in this group of individuals, and further evaluation should be performed if 1) anti-phase I IgG continue to rise and the anti-phase II IgG titers remain constant or decrease and 2) there is evidence of inflammatory disease [42]. Short-term treatment with trimethoprim-sulfamethoxazole in pregnant women infected with Q fever was not effective in preventing intrauterine fetal death [2]. Long-term treatment with trimethoprimsulfamethoxazole (≥ 5 weeks) in one study was associated with no occurrence of intrauterine fetal deaths, fewer obstetric complications (including decreased risk of placental infection), and decreased occurrence of chronic Q fever [2]. However, the author noted selection bias in the last 4 years of the 9-year study, as most persons not receiving long-term trimethoprim-sulfamethoxazole were individuals who received a Q 0701016F.doc 3-83 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever fever diagnosis just before or immediately after delivery. In an earlier report of 11 women infected during the first trimester of pregnancy, abortion did not occur in four women who received cotrimoxazole treatment throughout pregnancy, but was observed in 6 untreated women and in one woman who received only 3 weeks of co-trimoxazole [3]. Additionally, C. burnetii was detected in the placenta by culture and/or PCR in 4 women who did not receive long-term antibiotics, but in only 2 of the 4 women who received long-term antibiotics. Serological evidence of chronic infection in this cohort was observed in 12 of 17 women infected in the first six months of pregnancy. Nine individuals received long-term treatment for chronic Q fever (doxycycline and hydroxychloroquine for 18 months). • Doxycycline, 100 mg every 12 hours for 14 to 21 days will shorten the duration of the illness. Fever usually disappears within 1 or 2 days after treatment is initiated. Resistance to doxycycline is uncommon but has been reported. If doxycycline cannot be used, macrolides or quinolones may be considered as alternative therapy [27,39]. Chloramphenicol has been used to treat Q fever [43]. • Successful treatment of Q fever endocarditis is much more difficult and should involve consultation with an infectious disease specialist and cardiologist. Prolonged treatment with doxycycline given in combination with hydroxychloroquine (or with ofloxacin) has been successful and associated with death rates of approximately 5% (compared to 40% mortality from Q fever endocarditis in earlier reports) [31,39,44]. The effect of chloroquine is felt to be due to the increase in lysosomal pH, that allows the doxycline to be bactericidal against C. burnetii (the acidic environment of the C. burnetii infection site is felt to inhibit the bactericidal activity of doxycycline) [29]. The currently recommended duration of treatment for Q fever endocarditis has been 18 months to 3 years for doxycycline and hydroxychloroquine, at least 4 years for doxycycline and ofloxacin therapy, and perhaps life-long treatment if the treatment regimen does not include doxycycline [27,31,44]. Heart valve replacement may be required to achieve a cure [45]. • Other treatment regimens for chronic Q fever include doxycycline with ciprofloxacin, doxycycline with rifampin, ciprofloxacin with rifampin, and doxycycline with co-trimazole [29,39]. Environmental Decontamination and Infection Control The spore-like form of the organism is very resistant to heat, drying, and sunlight. Fomites contaminated by blood, urine, feces, and birth fluids can remain highly infectious for long periods. Decontamination can be accomplished with soap and water of 0.5% hypochlorite solution. Health care workers should follow standard precautions when taking care of patients with infections associated with C. burnetii. They include wearing appropriate personal protective equipment (PPE), including masks and eye protection, when generation of aerosols or splatter of bodily fluids is anticipated. However, person-to-person transmission of C. burnetii is rare. Patients exposed to aerosolized C. burnetii do not present a risk for secondary contamination or reaersolization of the organism. However, contaminated clothing may be a source of infection. Patients do not need to be placed in private rooms. References 1. Fenoller F, Fournier P-E, Carrieri MP, Habib G, Messana T, Raoult D: Risk factors and prevention of Q fever endocarditis, Clin Infect Dis 2001; 33:312–16. 2. Carcopino X, Raoult D, Bretelle F, Boubli L, Stein A: Managing Q fever during pregnancy: the benefits of long-term cotrimoxazole therapy, Clin Infect Dis 2007; 45:548–55. 0701016F.doc 3-84 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever 3. Raoult D, Fenollar F, Stein A: Q fever during pregnancy. Diagnosis, treatment, and follow-up, Arch Int Med 2002:162:701-4. 4. Franz DR, Jahrling PB, Friedlander AM, McClain DJ, Hoover DL, bryne WR, Ravlin JA, Christopher GW, Eitzen EM: Clinical recognition and management of patients exposed to biological warfare agents, JAMA 1997; 278(5):399–411. 5. Pike RM: Laboratory-associated infections: summary and analysis of 3,921 cases, Hlth Lab Sci 1976; 13(2):105–14. 6. Johnson JE, Kadull PJ: Laboratory-acquired Q fever: a report of fifty cases, Am J Med 1966; 41(3):391–403. 7. Rusnak JM, Kortepeter MG, Hawley RJ, Anderson AO, Boudreau E, Eitzen E: Risk of occupationally acquired illnesses from biological threat agents in unvaccinated laboratory workers, Biosecur Bioterr. 2004; 2(4):281–93. 8. Rusnak JM, Kortepeter MG, Aldis J, Boudreau E: Experience in the medical management of potential laboratory exposures to agents of bioterrorism on the basis of risk assessment at the United States Army Medical Research Institute of Infectious Diseases (USAMRIID). J Occup Env Med. 2004; 46(8):801–11. 9. Rusnak JM, Kortepeter MG, Hawley RJ, Boudreau E, Aldis J, Pittman PR: Management guidelines for laboratory exposures to agents of bioterrorism, J Occup Env Med 2004; 46(8):791–800. 10. Hornibrook JW, Nelson KR: An institutional outbreak of pneumonitis, Pub Health Rep 1940; 55(40):1936–54. 11. Huebner RJ: Report of an outbreak of Q fever at the National Institute of Health, Amer J Pub Health 1947; 37:431–40. 12. Oliphant JW, Gordon DA, Meis A, et al. Q fever in laundry workers presumably transmitted from contaminated clothing. Am J Hyg. 1949;49:76-86. 13. Graham CJ, Yamauchi T, Rountree P: Q fever in animal laboratory workers: an outbreak and its investigation, Am J Infect Control 1988;17:345-48. 14. Hall CJ, Richmond SJ, Caul EO, Pierce NH, Silver IA: Laboratory outbreak of Q fever acquired from sheep, Lancet 1982; 1(8279):1004–06. 15. Centers for Disease Control: Q fever at a university research center-California, MMWR Morb Mort Wkly Rep 1979; 28:333–34. 16. Maurin M, Raoult D: Q fever, Clin Microbiol Rev 1999;12:518-53. 17. Anonymous: Experimental Q fever in man, Br Med J 1950; 1:1000. 18. U.S. Department of Health and Human Services, Public Health Service, Centers for Disease Control and Prevention, National Institutes of Health: Biosafety in Microbiological and Biomedical Laboratories, 5th ed, pp 193–94. Washington DC, U.S. Government Printing Office, 2007. Retrieved December 7, 2007 from www.cdc.gov/OD/ohs/biosfty/bmbl5/bmbl5TOC.htm 19. Babudieri B: Q fever: a zoonoosis, Adv Vet Sci 1959; 5:81-183. 20. Eklund CM, Parker RR, Lackman DB: A case of Q fever probably contracted by exposure to ticks in nature, Public Health Rep 1947;62:1413-16. 21. Welsh HH, Lennette EH, Abinanti FR, Winn JF: Q fever in California. IV. Occurrence of “Coxiella burnetii” in the placenta of naturally infected sheep, Public Health Rept 1951; 66:1473-77. 22. Bell EJ, Parker RR, Stoenner HG: Q fever. Experimental Q fever in cattle, Am J Public Health 1949;39:478-84. 0701016F.doc 3-85 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever 23. Byrne WR: Q fever in Textbook of Military Medicine: Medical Aspects of Chemical and Biological Warfare, Part I, pp 523–37. Washington DC, Office of the Surgeon General at TMM Publications, Borden Institute, 1997. 24. Turck WP, Howitt G, Turnberg LA, Fox H, Longson M, Matthews MB, Das Gupta R: Chronic Q fever, Q J Med 1976; 45:193–217. 25. Fournier PE, Marrie TJ, Raoult D: Diagnosis of Q fever, J Clin Microbiol 1998; 36:1823-34. 26. Sawyer LA, Fishbein DB, McDade JE: Q fever in patients with hepatitis and pneumonia: results of laboratory-based surveillance in the United States, J Infect Dis 1988; 158:497-8. 27. CDC website. http://www.cdc.gov/ncidod/dvrd/qfever/index.htm (accessed November 16, 2010) 28. Dupont HT, Thirion X, Raoult D: Q fever serology: cutoff determination for microimmunofluorescence, Clin Diag Lab Immunol 1994; 1:189–96. 29. Raoult D, Marrie T: Q fever, Clin Inf Dis 1995; 20:489-96. 30. Raoult D, Casalta JP, Richet H, Khan M, Bernit E, Rovery C, Branger S, Gouriet F, Imbert G, Bothello E, Colart F, Habib G: Contribution of systemic serological testing in diagnosis of infective endocarditis, J Clin Microbiol 2005; 43:5238-42. 31. Raoult D, Abbara S, Jassal DS, Kradin RL: Case 5-2007: A 53-year-old man with a prosthetic aortic valve and recent onset of fatigue, dyspnea, weight loss, and sweats, NEJM 2007; 356:715-25. 32. Healy B, Llewelyn M, Westmoreland D, Lloyd G, Brown N: The value of follow-up after acute Q fever infection, J Infect 2006; 52:e109-12. 33. Peter O, Flepp M, Bestetti G, Nicolet J, Luthy R, Dupuis G: Q fever endocarditis: diagnostic approaches and monitoring of therapeutic effects, Clin Invest 1992; 70:932-37. 34. Guigno D, Couopland B, Smith EG, Farrell ID, Desselberger U, Caul EO: Primary humoral antibody response to Coxiella burnetii, the causative agent of Q fever, J Clin Microbiol 1992; 30:1958-67. 35. Waag D, Chulay J, Marrie T, England M, Williams J: Validation of an enzyme immunoassay for serogiagnosis of acute Q fever, Eur J Clin Microbiol Infect Dis 1995; 14:421-42. 36. Benenson AS, Tigertt WD: Studies on Q fever in man, Trans Assoc Am Phys 1956; 69:98–104. 37. Yeamen MR, Mitscher LA, Baca OG: In vitro susceptibility of Coxiella burnetii to antibiotics, including several quinolones, Antimicrob Agents Chemother 1987; 31(7):1079–84. 38. Sobradillo V, Zalacain R, Capebastegui A, Uresandi F, Corral J: Antibiotic treatment in pneumonia due to Q fever, Thorax 1992; 47:276–78. 39. Gilbert DN, Moellering RC, Eliopoulos GM, Chambers HF, Saag MS, Eds: The Sanford Guide to Antimicrobial Therapy 2009 (39th Edition). Antimicrobial Therapy, Inc, Sperryville, Virginia, 2009. 40. Kofteridis D, Gikas A, Spiradakis G, et al. Clinical response of Q fever infection to macrolides. Presented at the Fourth International Conference on Macrolides, Azalides, Streptogramins and Ketolides, Barcelona, Spain, January 12-23, 1998. 41. Kuzman I, Schonwald S, Culig J, et al. The efficacy of azithromycin in the treatment of Q fever: a retrospective study. Presented at the Fourth International Conference on Macrolides, Azalides, Streptogramins and Ketolides, Barcelona, Spain, January 12-23, 1998. 42. Armed Forces Infectious Diseases Society Triservice Q fever Working Group: Clinical practice guidelines for management of patients with Q fever. Bandera, Texas, April 13, 2008. 43. Pierce TH, Yucht SC, Gorin AB, Jordan GW, Tesluk H, Lillington GA: Q fever pneumonitis: diagnosis by transbronchoscopic lung biopsy, West J Med 1979; 130(5):453–55. 0701016F.doc 3-86 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever 44. Raoult D, Houpikian P, Tissot-Dupont H, Riss JM, Arditi-Djiane J, Brouqui P: Treatment of Q fever endocarditis: comparison of 2 regimens containing doxycycline and ofloxacin or hydroxychloroquine, Arch Inter Med 1999;159:167-73. 45. Landais, C, Fenollar F, Thuny F, Raoult D: From acute Q fever to endocarditis: serological follow-up strategy, Clin Infect Dis 2007; 44:1337-40. Table 3-16. Diagnostic Testing for Q Fever Specimen Site and Test Serum for PCR Serum for serological tests (IFA is gold standard, macroagglutination, microagglutination, ELISA) Method of Collection Transport Comments Whole blood is preferable (collected in citrate tube) for PCR; serum (spun from serum collected in red top or TT tube) or citrate plasma (spun from whole blood collected in citrate tube) is secondary choice of specimen. Collect a minimum of 1 mL (7 mL is adequate) of whole blood, serum, or citrate plasma for PCR. Transport chilled (not frozen). PCR is investigational. PCR specimens should be obtained in acute phase and prior to antibiotics. Collect 1 mL minimum (7 mL adequate) serum for serology in either red top or TT tubes and spun. Refrigerate at 2ºC to 4ºC for 5 to 7 days. Significant antibody titers are usually not identifiable until 2 to 3 weeks into the illness. Freeze at -30ºC or lower if greater than 7 days. Avoid multiple freeze–thaw cycles. Store in aliquots of at least 100 μL in O-ring screw-cap vials to avoid evaporation. Samples may be preserved by adding thimerosal to a final concentration of 0.01%. A four-fold rise in titers is considered diagnostic for Q fever. High titers to Phase II antigen occur in acute infection and high titers to Phase I antigen in chronic infection. IFA antiphase II IgM titer ≥ 1:40 or anti-phase II IgG ≥ 1:200 are used for criteria for diagnosis of acute Q fever. However, IgM titers by IFA have been reported to persist for a year or longer in 3% of persons with Q fever. IFA anti-phase I titer of ≥ 1:200 is considered diagnostic of chronic Q fever. A CF titer of ≥ 1:8 to Phase I or Phase II antigen is considered to be positive for Q fever infection. ELISA may be used for diagnosis of acute infection. 0701016F.doc 3-87 Chapter 3 – Part I. Bacterial and Rickettsial Agents/Diseases – Q Fever Table 3-16. Diagnostic Testing for Q Fever (cont.) Collect > 1 mL of a lower respiratory specimen into a sterile container for stains and culture. Transport at room temperature, when transport time is < 2 hours. For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. Impression smears can be stained with Giemsa or Gimenez stains or processed for direct immunofluorescence detection for C. burnetii. Smears should be made promptly if possible. A smear should not be too thick; slides should be dried well and fixed. Samples left at room temperature may develop heavy overgrowth contaminants. Clinical samples of affected tissues can be submitted as aspirates or tissue biopsies for use in IHC, PCR, or culture procedures. Collect samples in a sterile screw top container. Transport at room temperature, when transport time is < 2 hours. For transport times between 2 to 24 hours, transport at 2ºC to 8ºC. Impression smears can be stained with Giemsa or Gimenez stains or processed for direct immunofluorescence detection for C. burnetii. Sputum for DFA and PCR Tissue for DFA, PCR, immunofluorescent stain, or electron microscopy Smears should be made promptly if possible. A smear should not be too thick; slides should be dried well and fixed. Samples left at room temperature may develop heavy overgrowth contaminants. Caution: Clinical isolates should be handled under BSL-3. Culture (generally not obtained as requires animal inoculation or cell culture) Collect 10 mL sterile whole blood in a syringe and cap. Transport capped syringe with whole blood at room temperature. Caution: Clinical isolates should be handled under BSL-3. Refrigerate if transport time > 1 hour. Culture requires animal inoculation or cell culture. *G supplement to medium: Add 1%–5% glucose, 5% glycerol, or meat infusion nutrient agar. 0701016F.doc 3-88
