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Acknowledgements
I would like to thank the following persons and institutions:
• Dr. Namrita Lall, my supervisor, for her guidance, easily approachable, friendly,
support and comments. Also, Prof. J.J. Marion Meyer, for giving me an opportunity to
study in the Botany Department, for his kind assistance, and critical comments. Special
thanks for their contribution during the research project, thesis write up and article
publication.
• Ms. Elsa van Wyk for the identification of the plant species.
• Supportive staff of the Medical Research Council in Pretoria, for antituberculosis tests.
• Ms. Cathy and Ms. Andrea Prinsloo of the Department of Pathology for their guidance
and assistance with regard to antibacterial tests.
• Dr. Quenton Kritzinger for his assistance and willingness during antifungal tests.
• Mr. Frank van der Kooy, for his kind assistance with regard to extraction, isolation,
chromatography and spectroscopy techniques throughout the entire study.
• Mr. Eric Palmer, for kindly providing his technical assistance with NMR spectroscopy.
• Dr. Lynne Collette of Department of Chemistry for analyzing the spectra of isolated
compound.
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Acknowledgements
• Dr. Ahmed Hussein from Egypt for his kind assistance during the experimental
procedures, antimicrobial assays, isolation and analysing the spectra of isolated
compounds.
• The Research team of the Department of Botany, Plant Physiology, for their assistance
in plant collection.
• The University of Pretoria International Affairs funding for giving me an opportunity
and financial assistance to go to King’s College London.
• Prof. P.J. Houghton for giving me an opportunity to get familiar with different
techniques of isolation and his assistance in King’s College London. I wish to express my
sincere thanks to the Pharmacognosy Research team at King’s College London.
• The National Research Foundation (NRF), the Innovation Funding (Technology
Missions), the Medical Research Council (MRC) and the University of Pretoria for
financial support.
• My family, grandmother Monica; moms Dianah and Nelly; sisters and brother
Michelle, Monica, Hope, Sabina, Charity and Abner for their encouragement,
understanding, support, patience, kindness throughout my study.
• Friends who encouraged and guided me during my entire study period.
• I thank God for His blessings and protection everyday.
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Chapter 12
APPENDICES - Publications
12.1 Publications resulting from this thesis:
S.P.N. Mativandlela, N. Lall and J.J.M. Meyer. Antibacterial, antifungal and
antituberculosis activity of the roots of Pelargonium reniforme CURT and Pelargonium
sidoides DC. Submitted to South African Journal of Botany
12.2 Article in preparation:
S.P.N. Mativandlela, N. Lall, A.A.H. Huisen and J.J.M. Meyer. Antituberculosis activity
of compounds isolated from Pelargonium siodides DC.
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Antibacterial, antifungal and antituberculosis activity of the roots of Pelargonium
reniforme Curtis and Pelargonium sidoides DC.*
SPN Mativandlela, N Lall* and JJM Meyer
Department of Botany, University of Pretoria, Pretoria 0002, South Africa
* Corresponding author, e-mail: [email protected]
Abstract
Root extracts of Pelargonium reniforme Curtis and Pelargonium sidoides DC. were
evaluated against bacteria which cause bronchitis, three fungal pathogens and
Mycobacterium tuberculosis using the agar diffusion and BACTEC radiometric method,
at concentrations ranging from 5.0 to 0.5 mg/ml. The ethanol extract of the roots of P.
sidoides inhibited the growth of Haemophilus influenza, Moraxella catarrhalis and
Staphylococcus pneumonia at a concentration of 5.0 mg/ml. Both acetone and ethanol
extracts of P. reniforme and only the ethanol extract of P. sidoides inhibited the growth
of Aspergillus niger and Fusarium oxysporum significantly at a concentration of 0.5
mg/ml. Growth of Rhizopus stolonifer was suppressed by the ethanol extract of P.
reniforme and P. sidoides at 5.0 and 1.0 mg/ml, respectively. Acetone, chloroform and
ethanol extracts of P. reniforme showed activity against M. tuberculosis exhibiting a
minimum inhibitory concentration of 5.0 mg/ml.
Keywords: Pelargonium reniforme; P. sidoides; Antibacterial; Antifungal;
Antituberculosis
* Corresponding author. Tel.: +27 12 420 2524; fax: +27 12 362 5099
Email address: [email protected]
* Submitted to South African Journal of Botany
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Introduction
The importance of Pelargonium species (Geraniaceae) is well documented (Watt and
Breyer-Brandwijk 1962, Hutchings 1996). The genus Pelargonium comprises of more
than 250 natural species of perennial small shrubs, which are limited in their geographical
distribution. About 80% of Pelargonium species are confined to the southern parts of
Africa, while others occur in Australia, New Zealand and the far East. These species
usually grow in short grassland and sometimes with shrubs and trees on stony soil
varying from sand to clay-loam, shale or basalt. The plants are evergreen when
cultivated, but die back in nature during droughts and winter (May to August) (Van der
Walt and Vorster 1985).
P. reniforme Curtis and P. sidoides DC. are highly valued by traditional healers
for their curative properties and they are well known to generations of Khoi / San and
Xhosa (South African tribes) traditional healers (Wagner and Bladt 1975). The Xhosa
and the Zulu tribes of South Africa use these species to treat coughs, diarrhoea and
tuberculosis (Watt and Breyer-Brandwijk 1962). The medicinally active ingredients are
found in the bitter tasting roots of the plants (Helmstader 1996). A commonly used
medicine produced in Germany, named, ‘Umckaloabo’ originates from the roots of P.
sidoides and P. reniforme (Helmstader 1996, Kayser and Kolodziej 1998). This herbal
medicine is extensively used in Germany for bronchitis, antibacterial and antifungal
infections. Although this herbal medicine (Umckaloabo) is successfully employed in
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modern phytotherapy in Europe to cure infectious diseases of the respiratory tract, the
scientific basis of its remedial effect is still unclear (Kayser and Kolodziej 1995).
Bacteria, which are associated with either primary or secondary infections of bronchitis,
are Streptococcus pneumoniae, Haemophilus influenzae and Moraxella catarrhalis. H.
influenzae, a Gram-negative bacterium, is an obligate human parasite that is passed from
person to person by way of the respiratory route. M. catarrhalis, a Gram-negative
bacterium, causes bronchitis and pneumonia in children and adults. S. pneumoniae, a
Gram-positive bacterium, infects the upper respiratory tract and can cause pneumonia,
also it can infect the lining of the brain-spinal cord (meningitis), bones (osteomyelitis),
joints (arthritis), ears (otitis media) and sinuses (sinusitis and bronchitis),
(Benjamin et al. 1991).
Apergillus niger, Fusarium oxysporum and Rhizopus stolonifer are some of the
fungal pathogens that can affect the respiratory tract. A. niger, is a causative agent of
pulmonary diseases such as asperigillosis, bronchial asthma, acute allergic alveolitis etc.
The fungus colonizes old tuberculous or bronchiostatic cavities, in which it forms a large
colony (aspergilloma); or it may actually invades the lung tissue to produce haemorrhagic
and necrotozing pneumonia (MacSween and Whaley 1992). F. oxysporum is responsible
for fusariosis, skin infection, respiratory tract infections (tuberculosis and bronchitis) and
arthritis and produces a 76% mortality rate in hospitalised immunocompromised patients
(Monier et al. 1994). R. stolonifer causes mucorosis disease and it has been reported that
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exposure to large numbers of Rhizopus spores has reportedly caused respiratory
complications (Alexopoulos et al. 1996). Previously, researchers have reported
antimicrobial activity of extracts of Pelargoniums and their constituents against a few
bacterial (St. aureus, S. pneumoniae, E. coli, K. pneumoniae, P. mirabilis, P. aeruginosa
and H. influenzae) and fungal (M. canis, M. gypseum, A. fumigatus, M. racemosus, R.
nigricans) pathogens as well as opportunistic yeasts such as C. albicans, C. glabrata, C.
krusei and Cryptococcus species (Latté and Kolodziej 2000). Plant extracts of P.
reniforme and P. sidoides have not been tested against the fungal pathogens, A. niger, F.
oxysporum, R. stolonifer and the Gram-negative bacteria M. catarrhalis, which are
indirectly responsible for the secondary infections of bronchitis and tuberculosis. In the
present study, we have investigated their antimicrobial activity against the bacteria and
fungi mainly responsible for bronchitis. We have also confirmed the findings of other
researchers on the antibacterial activity of these species against S. pneumoniae and H.
influenzae.
Tuberculosis (TB) kills approximately 2 million people each year, the global
epidemic is growing and becoming more problematic. The breakdown in health services,
the spread of HIV / AIDS and the emergence of multidrug-resistant (MDR) TB are
contributing to the worsening impact of this disease. It is estimated that between 2002
and 2020, approximately a billion people will be newly infected, more than 150 million
people will get sick, and 36 million will die of TB. The current threat in TB treatment lies
in the emergence of strains resistant to two of the best antituberculosis drugs, isoniazid
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(INH) and rifampicin (RIF). The current TB-treatment comprises of 3-4 drugs for a
period of 6-9 months (Bloom 2002). Novel drugs are required which can shorten this
long treatment period and target multidrug resistant strains of TB. However, nothing had
been published on antituberculosis activity of these plants. This is the first report on
antituberculosis activity of these plants.
Materials and Methods
Plant material
Roots of P. reniforme and P. sidoides were collected from Qwaqwa, a region in the Free
State province of South Africa. Voucher specimens of P. reniforme (P 092558) and P.
sidoides (P 092559) were deposited and identified at the H.G.W.J. Schweickerdt
Herbarium (PRU), University of Pretoria, South Africa.
Preparation of extracts
Dried and powdered roots of P. reniforme and P. sidoides (300g each) were extracted
with 1L of acetone, chloroform and ethanol separately. The extracts were filtered and
concentrated with a rotary vacuum evaporator to dryness at reduced pressure. For
antibacterial and antifungal assays, acetone and ethanol extracts were dissolved in
acetone to a concentration of 50mg ml-1 and 100mg ml-1 respectively. For the
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antituberculosis assay, all 3 extracts were dissolved in dimethyl sulphoxide (DMSO) to a
concentration of 500mg ml-1.
Micro-organisms
Bacteria
The bacteria used in this investigation H. influenzae, M. catarrhalis, and S. pneumoniae
were obtained from the Department of Pathology, University of Pretoria, South Africa
and maintained on Colombian agar.
Antibacterial assay
For the antibacterial assay, the minimum inhibitory concentration (MIC) of the acetone
and ethanol extracts were determined by incorporating various amounts (5.0, 1.0 and
0.5mg ml-1) of the extracts into chocolate agar in sterile bottles and placed in a water bath
to prevent solidification, then withdrawn into petri dishes and left to solidify for four
hours. The bacterial colonies were transferred into the sterile screw capped round tubes to
which 5ml of the diluting fluid (saline) was added for achieving McFarland no.1 turbidity
standard. Each suspension was streaked on petri dishes containing the extracts and the
chocolate agar. The plates (three replications) were incubated at 25 oC for 24 hours and
antimicrobial activity was evaluated thereafter. Streptomycin sulphate added to chocolate
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agar to a final concentrations of 0.5, 0.01 and 0.05mg ml-1 served as positive controls and
three petri dishes containing only 500 µl acetone mixed with chocolate agar served as
negative controls.
Fungi
The fungal pathogens used in the study, A. niger, F. oxysporum and R. stolonifer were
obtained from the Department of Microbiology and Plant Pathology, University of
Pretoria, South Africa. Each fungus was maintained on Potato Dextrose Agar (PDA) at ±
25 oC.
Antifungal assay
For the antifungal assay, the required amount of acetone and ethanol extracts were added
to sterile PDA in 5ml petri dishes before congealing to yield final concentrations of 5.0,
1.0 and 0.5mg ml-1. PDA plates with acetone and fungi served as controls. Once the agar
had solidified, a 5mm plug of a seven-day old fungal culture was placed in the centre of
the petri dish containing the extract-amended and unamended PDA plates. The plates
were sealed with parafilm and placed in a 25 oC incubator. Fungal growth was measured
on two diametric lines after 3, 6 and 9 days of growth. Each treatment was replicated
three times. The results of six-day growth was statistically analysed using analysis of
variance (ANOVA) and comparison of means by Duncan’s Multiple Range Test. The
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value of bars within each fungus are a mean of three replicates and are significantly
different P < 0.01(Figure 1).
Mycobacterium tuberculosis
A drug-susceptible strain of M. tuberculosis, H37Rv obtained from American Type, MD,
USA Culture Collection (ATCC), 27294, was used to investigate the activity of the plant
extracts.
Antituberculosis assay
The radiometric respiratory technique using the BACTEC system was used for
susceptibility testing of M. tuberculosis as described previously (Lall et al. 2001, Lall et
al. 2003). Solutions of all the extracts were prepared in DMSO to obtain a concentration
of 500mg ml-1 and stored at 4oC until used. Subsequent dilutions were made in DMSO
and added to BACTEC 12B vials containing 4 ml of 7H12 medium broth to achieve the
desired final concentrations of 5.0, 2.5, 1.0 and 0.5mg ml-1 together with PANTA
(Becton Dickinson & Company, Ferndale, South Africa), an antimicrobial supplement.
Control experiments showed that the final amount of DMSO (1 %) in the medium
had no effect on the growth of M. tuberculosis. Anti-TB drugs; streptomycin (4µg ml-1),
isoniazid (2µg ml-1), rifampicin (0.2µg ml-1) and ethambutanol (6µg ml-1) (Sigma
Chemical Co., South Africa), were also tested against the H37Rv strain of M.
tuberculosis which served as positive drug controls. A homogenous culture (0.1ml) of all
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the strains of M. tuberculosis, yielding 1 x 104 to 1 x 105 colony-forming units per
millilitre (CFU ml-1), was inoculated in the vials containing the extracts as well as in the
control vials (Heifets et al. 1985). Three extract-free vials were used as controls (medium
+ 1% DMSO): two vials (V1) were inoculated in the same way as the vials containing the
extracts, and one (V2) was inoculated with a 1:100 dilution of the inoculum (1: 100
control) to produce an initial concentration representing 1% of the bacterial population (1
x 102 to 1 x 103CFU ml-1). The MIC was defined as the lowest concentration of the
extract that inhibited > 99 % of the bacterial population.
When mycobacterium grows in 7H12 medium containing 14 C-labelled substrate
(palmitic acid), they use the substrate and
14
CO2 is produced. The amount of
14
CO2
detected reflects the rate and amount of growth occurring in the sealed vial, and is
expressed in terms of the growth index (GI), (Middlebrook et al. 1977). Inoculated
bottles were incubated at 37oC and each bottle was assayed every day to measure GI, at
about the same hour(s) until cumulative results were interpretable. The difference in the
GI values of the last two days is designated as
GI. The GI readings of the vials
containing the test extracts were compared with the control vials (V2). Readings were
taken until the control vials, containing a 100 times lower dilution of the inoculum than
the test vials, reached a GI of 30 or more. If the GI values of the vials containing the test
extracts were less than the control vials, the population was reported to be susceptible to
the compound. Each test was replicated three times.
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Whenever results suggested contamination (e.g., large, rapid increase in GI), bottles were
inspected and the organisms were stained by Ziehl-Neelsen stain to determine whether
the visible microbial growth was a mycobacterium (Kleeberg et al. 1980). With this stain,
the bacilli appear as brilliantly stained red rods against a deep sky-blue background.
Organisms often have a beaded appearance because of their polyphosphate content and
unstained vacuoles (Joklik et al. 1968).
Since anecdotal evidence suggests the use of a combination of ethanol extracts of
two Pelargoniums, we also included the combination of ethanol extracts of both the roots
for antimicrobial assay.
Results and Discussion
It was found from the antibacterial assay that the ethanol and acetone extracts of P.
sidoides and its combination with P. reniforme was active at 5.0mg ml-1 against H.
influenzae, M. catarrhalis and S. pneumoniae. Activity of streptomycin sulphate was
observed on each bacteria at 0.5, 0.01 and 0.05mg ml-1. Kayser and Kolodziej (1997),
found inhibition of S. pneumoniae and H. influenzae at concentrations of 7.5 and 5.0mg
ml-1 respectively by acetone extracts of roots of P. reniforme. There have been few
reports of these bacterial organisms being susceptible to other plant extracts. Christoph et
al. 2001 found antibacterial activity of Australian tea tree oil from Melaleuca alternifolia
(Cheel) and niaouli oil isolated from M. quinquenervia at 0.01 (%v v-1) against M.
catarrhalis. Crude acetone extracts of P. reniforme was not active against these bacteria
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at the highest concentration (5.0mg ml-1) tested, similar to the findings of Magama et al.
2002, using Euclea crispa extremely. Essential oils of P. graveolens were found to be
inactive against Moraxella sp. (Lis-Balchin et al. 1998b). Gram-negative bacteria have
been found to be less susceptible to plant extracts in earlier studies done by other
researchers (Kuhnt et al. 1994; Afolayan and Meyer 1995). Various other species of
Pelargoniums such as P. tomentosum, P. Odaratissium, P. denticulatum and P. filcifolum
have been found to possess good antibacterial activity against Gram-positive bacterial
species such as Staphylococcus aureus, Proteus vulgaris, Bacillus cereus, and S.
epidermidis (Lis-Balchin et al., 1998a).
The antifungal assay of the acetone and ethanol root extracts of P. reniforme and
ethanol root extract of P. sidoides, showed activity against the fungal pathogens tested at
a concentration of 5.0mg ml-1 (Figure 1: a-b). In in vitro antifungal bioassays conducted
earlier, Latté and Kolodziej (2000), found that the aqueous acetone extracts of the roots
of P. reniforme were less potent exhibiting a MIC of 8.0mg ml-1 against the filamentous
fungi (Aspergillus fumigatus, Rhizopus nigricans, Penicillium italicum) mold fungi and
opportunistic yeasts tested. Other plant extracts have been found to be antifungal against
the fungi tested in this study. Chandrasekaran and Venkatesalu (2004), investigated the
water and methanol extracts of Syzygium jambolanum for antifungal activity against A.
niger and R. stolonifer and the highest zones of inhibition were recorded at 1.0 and 0.5mg
ml-1 respectively. Chamundeeswari et al. 2004 found an MIC of 2.5mg ml-1 when ethanol
root extract of Trewia polycarpa was tested against A. niger.
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The antituberculosis assay of extracts was interpreted on day five or six when the control
vials (V2) reached a GI value of 30 or more (Table 1). Acetone, chloroform and ethanol
extracts from the roots of P. reniforme showed inhibitory activity at 5.0mg ml-1 against
the drug-sensitive strain of M. tuberculosis but the combination of root extracts from both
Pelargoniums did not show any synergy. Activities of the standard antituberculosis drugs
(streptomycin, ethambutol, rifampicin and isoniazid), used as positive controls, were
much stronger than those of the extracts. Our results are in agreement with previous
reports on antituberculosis activity of water extracts of Thymus vulgaris, Nidorella
anomala, Cryptocarya latifolia and acetone extract of Rapanea melanophloeos where
MIC’s were also found to be 5.0mg ml-1 against M. tuberculosis (Lall and Meyer 1999).
Extracts of P. sidoides were not active against M. tuberculosis similar to the results
obtained by Fabry et al. 1998 where methanol extracts of Entada abyssinica, Terminalia
spinosa, Harrisonia abyssinica, Ximenia caffra, Azadirachta indica and Spilanthes
mauritiana were found to be inactive against M. tuberculosis at concentrations ranging
from 2.0 – 0.5mg ml-1.
The bacteria and fungi inhibited in this study have been associated with infections
of the respiratory tract, which the local inhabitants of South Africa treat using roots of P.
sidoides and P. reniforme. Also, the fact that the extracts inhibited H. influenzae, M.
catarrhalis and S. pneumoniae, which cause bronchitis, provides some scientific rationale
for the use of the extracts for tuberculosis, bronchitis and chronic asthma.
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Acknowledgements
Thanks to Dr Bernard Fourie and the technical assistants of the Medical Research
Council (Pretoria) and Mahdi Ziaratnia for their assistance. The National Research
Foundation supported the research financially.
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60
a
a
a
50
a
b
40
30
A.niger
a a
F.oxysporum
R.stolonifer
b
b bc
c
c
d d
20
d
d
c bc
c
d
10
e
Et
/5
Et
/1
.5
Et
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on
tr
ol
Et
/C
A
C
A
C
/5
/1
5
/0
.
A
C
A
C
/C
on
tr
ol
0
(a)
60
a
aa
50
a
a
40
a
b
b
a
ab
30
a a
a
b
c
A.niger
F.oxysporum
R.stolonifer
ab
ab
b
b
c
c
20
c
10
Et
/5
Et
/1
Et
/0
.5
tr
ol
Et
/C
on
/5
A
C
/1
A
C
A
C
/0
.5
A
C
/C
on
tr
o
l
0
(b)
Captions for figures
Figure 1: Antifungal activity of:
(i) P. reniforme acetone and ethanol extract
(ii) P. sidoides acetone and ethanol extract
The value of bars within each fungus are a mean of three replicates and are significantly
different P < 0.01. ∗ AC = acetone; ET = ethanol
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University of Pretoria etd – Mativandlela, S P N (2005)
Chapter 12
Appendices
Table 1: Antituberculosis activity of the root extracts against the sensitive strain (H37Rv)
of Mycobacterium tuberculosis as determined by the radiometric method
Plant species
Sensitive strain
MICa (mg ml-1)
GIb values (mg ml-1)
Pelargonium reniforme (acetone)
5.0
1.5 ± 0.7 (Sc)
P. reniforme (chloroform)
5.0
0.5 ± 0.7 (S)
P. reniforme (ethanol)
5.0
2.5 ± 0.7 (S)
P. reniforme + P. sidoides (acetone)
5.0
-1.0 ± 2.8 (S)
P. reniforme + P. sidoides (chloroform)
5.0
1.0 ± 0.0 (S)
P. reniforme + P. sidoides (ethanol)
5.0
1.5 ± 0.7 (S)
P. sidoides (acetone)
nad
35.5 ± 6.3 (Re)
P. sidoides (ethanol)
na
276.0 ± 9.89 (R)
P. sidoides (chloroform)
na
18.5 ± 4.94 (R)
Ethambutol
0.006
0.33 ± 0.0 (S)
Rifampicin
0.0002
0.0 ± 0.0 (S)
Isoniazid
0.002
4.0 ± 0.0 (S)
Streptomycin
a
Minimum inhibitory concentration.
b
c
GI value (mean ± SD) of the control vial was 20 ± 1.4 for the sensitive strain.
Susceptible.
d
Not active at highest concentration tested.
e
Resistant at the highest concentration tested.
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