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NURSING
Nursing critically ill patients in
the intensive care unit
Elle Haskey
Critically ill patients present a challenge to the whole veterinary team
because they require invasive diagnostic tests, advanced procedures
and intensive nursing care. It is important to start with an assessment of
the patient to prioritise concerns and develop a care plan tailored for the
individual. The nursing process involves the implementation and delivery of
this plan, followed by evaluation of the nursing/treatments. This is a cyclical
process that is repeated over and over again as the patient’s key parameters
are reassessed and a further plan is made. The aim of this article is to
explore some key aspects of critical care nursing, including infection control
and hand hygiene, the management of tubes and lines, and dealing with
recumbent patients.
Infection control and hand
hygiene
A health care-associated infection (HCAI) has
been defined by the World Health Organization
(WHO) in 2011 as: ‘an infection occurring in a
patient during the process of care in a hospital
or other health-care facility which was not
present or incubating at the time of admission.
This includes infections acquired in the
hospital, but appearing after discharge, and
also occupational infections among staff of the
facility.’ WHO estimated that the prevalence of
HCAIs in people in 2010 was 7.6 per cent, and
this increased to 30 per cent in patients in an
intensive care unit due to several factors:
■■
Invasive devices in situ;
■■
Immunosuppression;
■■
Trauma;
■■
Reduced nutritional intake;
■■
Fatal disease process.
While the prevalence of HCAIs in animals
is unknown, we can assume it is similar to the
level in people. It is imperative that measures
are implemented to minimise the risks to the
patient because the impact of an HCAI includes
resistant pathogens, financial burden, a longer
hospital stay and an increase in both morbidity
and mortality.
Elle Haskey,
Queen Mother Hospital for Animals, Royal
Veterinary College, Hawkshead Lane, North
Mymms, Hertfordshire AL9 7TA, UK
e-mail: [email protected]
Many studies highlight that one of the
most effective methods for reducing the
transmission of pathogens between patients
is effective hand hygiene. Veterinary clinics
should have a hand hygiene policy that draws
attention to:
■■
■■
■■
Hand care (actions that reduce skin
irritation or damage);
Hand decontamination (with both hand
wash and alcohol hand rub);
Surgical hand preparation;
■■ A
ppropriate
use of both sterile and nonsterile gloves.
The WHO and Centers for Disease Control
and Prevention (CDC) have both published
guidelines on hand hygiene (CDC 2002, WHO
2009).
The area around sinks should be kept
fully stocked and clean at all times. Hand
decontamination takes up to 60 seconds with a
hand wash and only 20 seconds with an alcohol
hand rub. Veterinary nurses should abide by the
WHO (2011) recommendation of ‘five moments
for hand hygiene’:
■■
Before touching a patient;
■■
Before a clean/aseptic procedure;
■■ A
fter
body fluid exposure risk;
■■ A
fter
touching a patient;
■■ A
fter
touching a patient’s surroundings.
According to WHO (2013), adherence
to hand hygiene policies is around 45 to 65
per cent, with reasons for poor compliance
including inadequate staffing levels (too
busy), forgetfulness, poor education and no
role models. Audits should be carried out to
highlight areas for further training to improve
adherence and help reduce the risk to patients
of HCAI.
Managing tubes and lines
Many critical patients will have multiple lines,
tubes or drains in situ during the management of
their disease. As mentioned previously, invasive
devices increase the risk of an HCAI, so it is
important to manage such devices appropriately
and have policies in place so that all staff adhere
to the same management protocol.
Intravenous catheters
Critical patients require venous access for the
administration of fluid therapy, medications and
blood products, which involves the placement
of intravenous (IV) catheters. These tubes can
also be used for haemodynamic monitoring and
haemodialysis.
A short, wide-bore catheter is often placed
in a peripheral vein in the emergency situation
Fig 1: Placement of a short-term, over-the-needle, peripheral intravenous catheter
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carried out as per a shortterm catheter. There is
no recommendation to
change central venous
catheters
routinely.
De-instrumenting should
occur when the catheter
is no longer required or
a peripheral IV catheter
can be placed as it is
associated with a lower
risk of complication (CDC
2011).
Arterial catheters
Fig 2: Triple lumen medial saphenous long-stay
line in a cat
for prompt access (Fig 1). These catheters are
readily available, simple to place and cheap.
Long-stay lines may be preferable in critical
patients because they provide some additional
benefits. They:
■■
■■
Cause less thrombophlebitis (because they
are made from materials that induce less
reaction) (CDC 2011);
Allow blood sampling;
■■ A
llow
multiple drugs to be administered at
the same time through separate lumens;
■■
Allow central venous pressure monitoring.
Arterial catheters are
extremely
useful
in
critical patients, allowing
the monitoring of arterial
Fig 3: Arterial line in the dorsopedal artery of a postoperative
blood
pressure
and
patient. The line was connected to a transducer for invasive blood
pressure measurement. The tape is clearly labelled with the word
sampling of arterial blood
‘arterial’ to prevent drugs being administered into the artery
gas. The placement of
arterial catheters takes
practice and can be
spillage onto the patient’s leg during the
very difficult in conscious and small patients
procedure should be cleaned as thoroughly as
(particularly cats). They are placed and
possible.
managed in a similar manner to short-term
Once placed, the catheter should be
IV catheters, with the only difference being
covered with a sterile dressing to keep it clean
that they are prone to occlusion and should be
and prevent patient interference. The dressing
flushed with saline hourly; alternatively, there
should be removed daily to assess the leg
are systems for monitoring blood pressure that
for signs of infection or inflammation. If the
allow continuous flushing.
catheter is not being used for IV fluids, it should
Patients with arterial catheters should
be flushed every four to eight hours. There is
ideally be monitored closely because
no evidence that heparinised saline is superior
detachment of the bung can result in rapid
to saline for maintaining catheter patency
blood loss. The lines should be clearly labelled
(Ueda and others 2013). There is, however,
so that no drugs are administered into the
evidence that catheter-related complications
artery (Fig 3), and removed when no longer
are associated with increased duration of
required because they cause limb ischaemia in
IV catheter placement (Seguela and Pages
cats (Mazzaferro 2009, Bowlt and others 2013).
2011, Parkes 2015). It is therefore sensible to
Tracheostomy tubes
consider replacing IV catheters after 72 hours.
Tracheostomy tubes (Fig 4) are used to bypass
Long-stay lines should be placed aseptically
the upper airway in patients with an airway
with skin preparation as for the placement
obstruction (eg, as a result of laryngeal
of short-term catheters. A sterile dressing
oedema, masses or trauma). Patients with
should be applied over the insertion site once
these tubes must be supervised constantly
in situ. Transparent dressings are ideal as it
because tube occlusion will cause asphyxiation.
is possible to view the site without having to
change the dressing each time. Alternatively, a
sterile gauze dressing can be used.
These lines should be checked daily in
a sterile manner and observed for signs of
infection or inflammation. Flushing should be
The tubes are usually placed centrally (eg,
in the jugular vein), but can also be placed in
larger peripheral vessels (eg, medial/lateral
saphenous vein) (Fig 2). Contraindications
include coagulopathies; jugular placement
should be avoided if there is a concern about
increased intracranial pressure.
There are several strategies that can reduce
the risk of complication during the placement
and management of IV
catheters.
Irrespective
of the site for short-term
catheter
placement,
hair should be removed
with a wide clip to avoid
contamination,
and
the skin then prepared
with a skin disinfectant
(ie, chlorhexidine) and
alcohol before placement.
Hand hygiene should be
carried out by the person
placing
the
catheter.
Consumables should be
prepared and contained
on a clean surface (not
Fig 4: Tracheostomy tubes. (a) Obturator, (b) closed lumen
the floor) before starting
inner cannula, (c) twist–lock lumen inner cannula, (d) cuffless
the procedure. Any blood
tracheostomy tube
26
Fig 5: Canine patient with a tracheostomy tube.
The stay sutures aid placement
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Fig 7: Closed suction drain in a dog with septic peritonitis
Fig 6: Dog with a chest drain after a thoracotomy. There is a light dressing at
the insertion site, a clamp and one-way valve to achieve occlusion, and a string
vest to prevent interference
A sterile replacement tube should be readily
available at all times (preferably next to the
patient’s kennel), together with a calculated
dose of anaesthetic agent for an emergency
situation. Patients should be preoxygenated for
about two minutes before the tube is changed/
removed/suctioned.
Tubes with inner cannulae are desirable as
these enable easy cleaning. The inner cannula
is removed by gently twisting to unlock it (with
clean, washed hands and disposable gloves),
placed under running water to remove debris/
secretions and then rinsed in sterile saline. If
it is encrusted, special tracheostomy brushes
can be used for cleaning. Do not use a bottle
brush or cotton wool buds to clean the inside of
the tube as this will lead to abrasions that allow
debris and secretions to adhere to the surface.
Tubes for smaller patients do not have
inner cannulae because their presence would
excessively reduce the inner lumen of the
tube. Therefore, these types of tubes should
be replaced every 24 hours after a period of
preoxygenation (Fudge 2009).
Nebulisation and humidification can help
prevent the airway and secretions from drying,
and can be carried out every four to eight hours.
Suctioning of the airway is recommended
in many texts and may be useful to remove
excessive secretions. This should be performed
with care as it may cause mucosal injury and
actually increase secretion production. A
sterile suction catheter is inserted into the
tracheostomy tube in an aseptic manner with
sterile gloves to the estimated level of the
carina (tracheal bifurcation). The catheter
width should be no more than 50 per cent of
the width of the internal diameter of the inner
cannula of the tracheostomy tube. A new
sterile suction catheter should be used for each
suctioning attempt. This procedure should be
abandoned if the patient becomes distressed.
When tracheostomy tubes are placed in
cats, shredded paper should be used instead
of normal cat litter due to the risk of inhalation
through the tracheostomy tube. Harnesses
should be used instead of neck leads for dogs.
There should be stay sutures present around
the tracheal rings dorsal and ventral to the
tracheal incision, allowing rapid ‘opening’ of
the tracheostomy site and replacement of the
tube (Fig 5).
Chest drains
Thoracostomy tubes (chest tubes) are used to
drain air or fluid from the pleural space and, as
for animals with tracheostomy tubes, constant
monitoring of patients with these tubes is
required. Indications for chest drain placement
include:
■■
■■
■■
Post-thoracotomy;
Medical management of a pleural effusion
(chylothorax or pyothorax);
Pneumothorax (ongoing air leakage);
■■ T
ension
■■
pneumothorax;
Penetrating chest injury.
All chest drains should be handled with
strict asepsis due the increased risk of HCAI
(Sigrist 2009). The drain insertion site should
be dressed with a light sterile dressing and
inspected twice daily for signs of infection,
inflammation, migration or leakage. Patients
should wear a vest made from a stretchy
material (Surgifix) or T-shirt to protect the tube
and prevent it from being accidentally removed,
as well as an Elizabethan collar to prevent
interference. All drains should be clamped
and/or occluded with a three-way tap/one-way
valve (Fig 6).
The frequency of draining depends on
the underlying disease process and severity.
Continuous suction may be required in some
circumstances, whereas intermittent draining
is sufficient in others. When manual draining is
performed, sterile gloves should be worn. All
fluid or air produced should be recorded. If the
patient is showing signs of respiratory distress,
aspiration of the chest tube should be attempted.
Drains should be removed when they are
no longer required or when the volume of air
or fluid produced is less than 2 ml/kg/24 hours
(Sigrist 2009).
Closed suction drains
There are many different closed suction
drains available. They are commonly used
for abdominal drainage after a contaminated
surgery (ie, septic peritonitis) but can also be
placed in contaminated wounds or wounds
with a large dead space. The skin insertion site
should be separate from the surgical wound,
should be dressed with a light sterile dressing
and inspected in a sterile manner every 24
hours (or more frequently if the dressing is
struck through). These drains are often bulky
and need to be attached to the patient to allow
mobility. This can be achieved using a couple
of butterfly wings on the tubing attached to the
skin, or tubular netting can be helpful (Fig 7).
Drains should be emptied wearing sterile
gloves and the frequency depends on the drain
and its function. They should always be emptied
if full or if negative pressure has been lost.
As for chest drains, closed suction drains
should be removed when they are no longer
required or when the volume produced is less
than 2 ml/kg/24 hours (Szabo and others 2011).
Urethral catheters
Indications for urethral catheterisation are to:
■■
Obtain a urine sample;
■■ P
erform
■■ R
elieve
■■
a radiographic contrast study;
a urethral obstruction;
Measure urine output;
■■ I mprove
nursing care in a critical patient or
after surgery to the lower urinary tract.
Urethral catheters are sometimes placed
intermittently but are more often indwelling
and the duration of catheterisation will depend
on the reason for placement. The catheters are
made from a variety of materials with varying
stiffness, tissue reactivity and minimisation of
biofilm formation. Foley catheters are chosen
for indwelling catheter placement because
the balloon at the end of the catheter can be
inflated to hold the catheter in place.
Urinary tract infections (UTIs) are the
most common HCAI in human hospitals.
In the veterinary literature, Smarick (2009)
reported that 10 to 52 per cent of cats and dogs
develop UTIs. Prophylactic antibiosis is not
recommended as this can lead to the growth
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Fig 8: Urinary catheter with a closed collection system. The
collection bag is double bagged to prevent contamination
and is anchored to the patient’s leg to reduce tension on the
urinary tract if the patient moves around
of resistant bacteria (Ogeer-Gyles and others
2006). The best way to reduce the likelihood
of a catheter-associated UTI is to follow good
hygiene principles. The CDC has published
guidelines for urinary catheter placement and
management (CDC 2009).
All urinary catheters should be placed in
a sterile manner (including the use of sterile
gloves), whether they are placed intermittently
or indwelling. Hair should be clipped from the
preputial/vulval area, with care taken not to
cause trauma to the skin as this may increase
the likelihood of an infection. The skin should
be prepared with a chlorhexidine or iodine skin
disinfectant. An appropriately sized catheter
should be selected for the patient – larger
catheters may cause increased irritation
to the urinary tract, thus increasing the
risk of infection. Lubrication of the catheter
can prevent trauma to the urethra and aid
placement. A closed urine collection system
should be attached to any indwelling urinary
catheter so that urine output can be quantified
and the risk of a UTI is reduced (Ogeer-Gyles
and others 2006, CDC 2009, Smarick 2009). In
addition, a closed system increases patient
comfort by preventing leakage and urine
scald. Urinary catheters can be anchored to
the patient’s tail or leg with tape to prevent
constant pulling on the catheter and trauma to
the bladder/urethra (Fig 8).
There is no veterinary study describing the
risk of UTI with/without cleaning the prepuce/
Fig 9: Faecal management system for a dog with parvovirus
vulva while the patient is catheterised.
However, there are recommendations to wipe
the outside surface of the catheter (proximal
to distal) with chlorhexidine swabs and to
flush the prepuce/vulva at regular intervals
(Smarick 2009).
The collection bag should be kept below
the level of the patient to prevent reflux and an
unobstructed urine flow should be maintained
at all times. Bags can be emptied as necessary
by disinfecting the drainage port before/
after drainage to minimise the number of
microorganisms entering the collection bag.
Routine flushing of the urinary catheter
is unnecessary and will increase the risk of
infection (Hooton and others 2010) so should
only be considered if there is an obstruction
in flow and should be carried out in a sterile
manner.
Recumbent patient
management
Recumbent patients are at risk of developing
pressure sores (decubital ulcers) and urine
and faecal scalding. The placement of a urinary
catheter may be contraindicated in some
situations, so patients need to be managed
carefully to reduce the risk of urine scald.
Ideally, patients should be kept clean and dry
at all times on bedding that wicks fluid away
(eg, Vetbed). If a patient becomes soiled, a
mild unmedicated shampoo should be used
to cleanse the soiled
areas; it is important
that hair and skin
are thoroughly dried.
This can be hugely
time
consuming
for veterinary staff
as bathing may be
required on a frequent
basis until the patient
is ambulatory. There
is little evidence to
support the use of
petroleum jelly or
zinc
oxide
cream
for managing scald,
Fig 10: Ocular care involves flushing the eyes with saline
although cornstarch
28
powder can be a useful adjunct as it helps to
dry the patient after bathing.
A faecal management system can be
considered for recumbent patients with profuse
diarrhoea (Fig 9) (MILA International 2014). It is
inserted into the rectum and inflated (similar to
a Foley urinary catheter) with a built-in closed
collection system. In my experience, it is well
tolerated and a time-saving tool when caring
for a recumbent patient with diarrhoea.
Frequently moving a recumbent patient
can help to minimise the risk of pressure
sores developing. Ideally, the animal should be
turned every two to four hours, with its front
end propped up to allow both lungs to inflate.
Foam wedges, pillows and duvets are useful
for padding beds. A passive and active range
of movement routines and massage can keep
patients supple and promote blood/lymph flow.
Hoists are useful for large patients so that they
can be stood up and encouraged to walk without
putting staff at risk of injury. Once patients
are ambulatory, physiotherapy sessions can
be extended to include sling walks, balancing
exercises and even hydrotherapy.
Ocular and oral care
Many critical patients have reduced
tear production and reduced blink. This
predisposes to corneal ulcer formation and
eye infections. Cats receiving high doses of
analgesia such as ketamine should have their
eyes lubricated to prevent drying and corneal
ulcer formation (Plumb 2008). Ulceration
can be avoided by applying sterile waterbased eye lubrication every two to four hours
(Clare 2009). If it is suspected that a patient
is developing a corneal injury, a fluorescein
stain of the cornea should be performed. Eyes
can be flushed with saline once to twice daily
(Fig 10) and if there are concerns with regards
to infection, topical antibiotics should be
administered.
Mechanically ventilated patients must also
receive oral care. These patients may not be
able to swallow and are at risk of developing
pneumonia as a result of oral bacterial
contamination of the lower airways. Hospitalacquired pneumonia is 20 times more likely to
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occur in mechanically ventilated patients than
others (Nakamura and Tompkins 2012).
Oral care should be performed wearing
gloves to minimise the transfer of bacteria
onto the hands of the nurse. Suctioning of
the mouth and pharynx may be beneficial if
there are copious oral secretions but this may
not be possible in conscious animals. Dilute
chlorhexidine mouthwash (Corsodyl; GSK)
(diluted 1:1) can be used to clean the mouth and
wipe around the teeth, helping to reduce the
bacterial load (Clare 2009). Care must be taken
in neurologically impaired patients to prevent
the nurse from being bitten. The tongue should
always be kept inside the mouth to prevent it
from drying or becoming swollen.
‘Big little things’
Nurses should work at creating a patient bond
by providing ‘TLC’ and grooming. The area of
the hospital where the critical patients are
housed should be quiet, with minimal traffic.
There should be the ability to dim lights to
mimic day/night and allow periods of quiet
time and rest. Sleep is an important medicine
that is often forgotten in veterinary hospitals.
Treatments should be appropriately grouped
to allow periods of rest and avoid disturbing
the patient unnecessarily. Noisy/stressed
animals should have measures taken to help
them settle and prevent them from upsetting
others (eg, quiet music can sometimes help). It
is amazing what can be achieved with anxious
patients if you just take things slowly and talk
to them. If there is time to bath/groom a patient
or transport it outside on a trolley to get some
fresh air this can help to motivate it to do a little
more. In some situations, it can be beneficial
to arrange owner visits, particularly in
‘depressed’ patients or perhaps those that are
not eating. However, if this stresses the patient,
it may be wise to avoid future visits.
As nurses are often busy and stretched, the
‘big little things’ are lower on the priority list.
Try to remember how you would want somebody
to treat your pet if it was in the hospital.
The critical cat
Cats behave differently from dogs in the
hospital environment; their response to disease
processes is often different from their canine
counterparts and being generally much smaller
creates different challenges in their management
for nurses. If there are some nurses within the
team who have that magic cat-whispering talent,
make sure they are used more for nursing feline
patients. Other nurses who are less confident
can then work alongside them to learn and
improve their cat knowledge and handling skills.
Provisions should be made to reduce stress
in hospitalised feline patients. The International
Society of Feline Medicine has published
recommended guidelines on handling and
nursing feline patients (Rodan and others 2011,
Carney and others 2012).
Summary
Nursing critically ill patients can sometimes
be overwhelming because there is so much
to think about. Nursing frameworks (such
as Kirby’s Rule of 20) can be useful to give
direction and enhance efficacy and efficiency
(Barton 2009). Policies and protocols can
be developed to ensure there is consistency
between staff members and to reduce the
risk of HCAIs. It can also be demanding and
frustrating, requiring input from the whole
veterinary team. Clear communication and
teamwork can lead to positive outcomes and
patients making a recovery, which is hugely
rewarding for all involved.
I am happy to be contacted to discuss
specific questions about aspects of critical
care nursing related to the article.
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doi: 10.1136/inp.i6051
Elle Haskey graduated from the University of Bristol in 2008 with an honours degree in veterinary
nursing and practice administration. She worked in a mixed practice in the Midlands before
returning to the University of Bristol as senior ICU nurse. She gained the AVECCT Veterinary
Technician Specialist (Emergency and Critical Care)qualification in 2012, and, in 2013, moved
to join the Royal Veterinary College ECC team where she is now head ECC nurse.
In Practice FOCUS November 2016
25-29 Haskey.indd 29
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27/10/2016 16:32
Downloaded from http://inpractice.bmj.com/ on May 2, 2017 - Published by group.bmj.com
Nursing critically ill patients in the intensive
care unit
Elle Haskey
In Practice 2016 38: 25-29
doi: 10.1136/inp.i6051
Updated information and services can be found at:
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