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Abstract

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The volatile oils of black pepper [Piper nigrum L. (Piperaceae)], clove [Syzygium aromaticum (L.) Merr. & Perry (Myrtaceae)], geranium [Pelargonium graveolens L'Herit (Geraniaceae)], nutmeg [Myristica fragrans Houtt. (Myristicaceae), oregano [Origanum vulgare ssp. hirtum (Link) Letsw. (Lamiaceae)] and thyme [Thymus vulgaris L. (Lamiaceae)] were assessed for antibacterial activity against 25 different genera of bacteria. These included animal and plant pathogens, food poisoning and spoilage bacteria. The volatile oils exhibited considerable inhibitory effects against all the organisms under test while their major components demonstrated various degrees of growth inhibition.

The antiseptic qualities of aromatic and medicinal plants and their extracts have been recognized since antiquity, while attempts to characterize these properties in the laboratory date back to the early 1900s ( Martindale 1910; Hoffman & Evans 1911). Plant volatile oils are generally isolated from nonwoody plant material by distillation methods, usually steam or hydrodistillation, and are variable mixtures of principally terpenoids, specifically monoterpenes [C10] and sesquiterpenes [C15] although diterpenes [C20] may also be present, and a variety of low molecular weight aliphatic hydrocarbons (linear, ramified, saturated and unsaturated), acids, alcohols, aldehydes, acyclic esters or lactones and exceptionally nitrogen- and sulphur-containing compounds, coumarins and homologues of phenylpropanoids. Terpenes are amongst the chemicals responsible for the medicinal, culinary and fragrant uses of aromatic and medicinal plants. Most terpenes are derived from the condensation of branched five-carbon isoprene units and are categorized according to the number of these units present in the carbon skeleton ( Dorman 1999).

The antimicrobial properties of plant volatile oils and their constituents from a wide variety of plants have been assessed ( Lis-Balchin & Deans 1997) and reviewed ( Janssen et al. 1987 ; Jain & Kar 1971; Inouye et al. 1983 ; Garg & Dengre 1986; Ríos et al. 1987 ; Sherif et al. 1987 ; Deans & Svoboda 1988, 1989; Cruz et al. 1989 ; Recio et al. 1989 ; Crespo et al. 1990 ; Carson et al. 1995 ; Larrondo et al. 1995 ; Pattnaik et al. 1995 ; Carson et al. 1996 ; Nenoff et al. 1996 ; Ríos et al. 1988 ). It is clear from these studies that these plant secondary metabolites have potential in medical procedures and applications in the cosmetic, food ( Ueda et al. 1982 ; Shelef 1983; Jay & Rivers 1984; Gallardo et al. 1987 ; Baratta et al. 1998a,b ; Youdim et al. 1999 ) and pharmaceutical industries ( Janssen et al. 1988 ; Pélissier et al. 1994 ; Shapiro et al. 1994 ; Cai & Wu 1996).

Investigations into the antimicrobial activities, mode of action and potential uses of plant volatile oils have regained momentum. There appears to be a revival in the use of traditional approaches to protecting livestock and food from disease, pests and spoilage in industrial countries. This is especially true in regard to plant volatile oils and their antimicrobial evaluation, as can be seen from the comprehensive range of organisms against which volatile oils have been tested. These have included food spoiling organisms ( Zaika et al. 1983 , 1984b; Connor & Beuchat 1984a; Janssen et al. 1988 ; Ouattara et al. 1997 ) and food poisoning organisms ( Beuchat 1976; Tharib et al. 1983 ; Deans & Ritchie 1987; Lis-Balchin & Deans 1997), spoilage and mycotoxigenic filamentous fungi ( Knobloch et al. 1989 ), pathogenic and dimorphic yeasts ( Boonchild & Flegel 1982; Ghannoum 1988) and animal and plant viruses ( Ieven et al. 1982 ; Romerio et al. 1989 ).

The aims of the present investigation were to assess the antimicrobial activities of the test volatile oils and compare these to the effect of the antibiotics upon bacterial growth; to assess the components determined to be present in the volatile oils where available; to use these data to deduce which components are likely to contribute to the activities of the whole oils and to determine any structural relationships between the components and their antibacterial activity.

MATERIALS and METHODS

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The volatile oils of black pepper [Piper nigrum L. (Piperaceae)], clove [Syzygium aromaticum (L.) Merr. & Perry (Myrtaceae)], geranium [Pelargonium graveolens L'Herit (Geraniaceae)], nutmeg [Myristica fragrans Houtt. (Myristicaceae)], oregano [Origanum vulgare ssp. hirtum (Link) Letsw. (Lamiaceae)] and thyme [Thymus vulgaris L. (Lamiaceae)] were screened for antimicrobial activity using an agar diffusion technique ( Deans & Ritchie 1987) against 25 microorganisms of significant importance ( Table 1). In addition, 21 authentic terpenoids and the phenylpropanoid eugenol, commonly found in these volatile oils, were also screened for activity ( Table 2).

Table 1.  Zones of growth inhibition (mm) showing antibacterial activity for a number of selected plant volatile oils; well diameter 4.0 mm
Bacterial strainSourceMyristica fragransOriganum vulgarePelargonium graveolensPiper nigrumSyzygium aromaticumThymus vulgaris
  1. Source of bacterial strains: NCIB, National Collection of Industrial Bacteria; NCTC, National Collection of Type Cultures; ATCC, American Type Culture Collection; NCPPB, National Collection of Plant Pathogenic Bacteria; NCDO, National Collection of Dairy Organisms. Values for zone of growth inhibition are presented as mean ± SEM.

Acinetobacter calcoaceticaNCIB 8250 12.7 ± 1.3 52.2 ± 1.513.0 ± 0.3 12.3 ± 2.010.3 ± 0.2 30.7 ± 0.5
Aeromonas hydrophilaNCTC 8049>90.0>90.0No inhibition>90.011.7 ± 1.1>90.0
Alcaligenes faecalisNCIB 81569.0 ± 0.3 33.6 ± 0.1No inhibition7.1 ± 0.823.1 ± 0.6 53.8 ± 1.2
Bacillus subtilisNCIB 36107.0 ± 0.4 20.5 ± 0.411.4 ± 0.69.5 ± 0.621.1 ± 0.1 23.4 ± 1.2
Beneckea natriegensATCC 14048 10.0 ± 1.2 37.1 ± 3.211.0 ± 0.7 10.8 ± 0.715.8 ± 0.7>90.0
Brevibacterium linensNCIB 8456 22.2 ± 0.3>90.07.6 ± 0.1 15.9 ± 1.029.8 ± 0.1>90.0
Brocothrix thermosphactaSausage meat9.7 ± 0.7 31.2 ± 0.88.6 ± 0.57.2 ± 0.111.1 ± 0.1>90.0
Citrobacter freundiiNCIB 11490 12.8 ± 0.1 29.6 ± 0.816.0 ± 2.0 12.0 ± 1.614.1 ± 2.6>90.0
Clostridium sporogenesNCIB 10696No inhibition>90.07.8 ± 0.68.7 ± 0.313.4 ± 0.5>90.0
Enterococcus faecalisNCTC 775 18.5 ± 1.2 17.9 ± 0.819.8 ± 2.18.8 ± 0.915.5 ± 0.6 41.8 ± 0.8
Enterobacter aerogenesNCTC 10006No inhibition 14.6 ± 0.1No inhibitionNo inhibition7.8 ± 1.1 15.2 ± 0.7
Erwinia carotovoraNCPPB 312 14.1 ± 2.6 31.2 ± 1.4No inhibition 12.9 ± 1.011.7 ± 0.4 35.8 ± 4.4
Escherichia coliNCIB 8879 10.4 ± 0.1 29.5 ± 3.4No inhibition7.3 ± 0.413.6 ± 0.3 32.4 ± 0.1
Flavobacterium suaveolensNCIB 8992 16.9 ± 0.99.4 ± 0.730.9 ± 5.4 10.1 ± 0.114.4 ± 0.2>90.0
Klebsiella pneumoniaeNCIB 418 16.9 ± 0.9 19.0 ± 1.513.8 ± 0.2No inhibition9.1 ± 0.1 31.8 ± 0.5
Lactobacillus plantarumNCDO 343No inhibition 23.8 ± 0.3No inhibitionNo inhibition28.5 ± 1.0 26.3 ± 0.4
Leuconostoc cremorisNCDO 543No inhibition>90.016.7 ± 2.3 16.3 ± 0.818.7 ± 0.6>90.0
Micrococcus luteusNCIB 8165 11.7 ± 0.3 21.5 ± 0.113.3 ± 0.4 12.4 ± 0.114.8 ± 0.8>90.0
Moraxella sp.NCIB 107626.4 ± 0.2 31.4 ± 1.9No inhibition5.4 ± 0.215.8 ± 0.8 29.0 ± 5.6
Proteus vulgarisNCIB 4175 10.0 ± 1.1 44.6 ± 4.9No inhibition7.1 ± 0.39.1 ± 0.6>90.0
Pseudomonas aeruginosaNCIB 950No inhibition>90.019.4 ± 0.17.7 ± 0.914.0 ± 1.9 33.5 ± 2.0
Salmonella pullorumNCTC 107048.4 ± 0.5 46.0 ± 6.76.9 ± 0.67.1 ± 0.214.0 ± 0.8>90.0
Serratia marcescensNCIB 13778.2 ± 0.3 18.9 ± 0.48.5 ± 0.47.5 ± 0.421.6 ± 0.9 39.1 ± 0.8
Staphylococcus aureusNCIB 6571 24.6 ± 0.4 17.6 ± 0.513.6 ± 0.3 14.5 ± 0.314.9 ± 0.1>90.0
Yersinia enterocoliticaNCTC 104607.3 ± 0.4 33.9 ± 0.4No inhibition 11.7 ± 2.213.7 ± 0.1 25.5 ± 2.9
Table 2a.  Zones of growth inhibition (mm) showing antibacterial activity for a number of selected plant volatile oil components; well diameter 4.0 mm
Bacterial strain1234567
  1. Values for zone of growth inhibition are presented as mean ± SEM. 1, Borneol; 2, δ-3-carene; 3, carvacrol; 4, carvacrol methyl ester; 5, cis/trans citral; 6, eugenol; 7, geraniol.

Acinetobacter calcoacetica7.0 ± 1.110.0 ± 0.245.3 ± 1.3No inhibition7.9 ± 0.915.4 ± 0.36.1 ± 0.2
Aeromonas hydrophila8.2 ± 0.511.1 ± 0.837.7 ± 2.4No inhibition7.9 ± 0.317.0 ± 0.46.4 ± 0.5
Alcaligenes faecalisNo inhibition14.4 ± 0.721.8 ± 1.65.9 ± 0.78.4 ± 0.412.3 ± 0.57.0 ± 0.2
Bacillus subtilis10.4 ± 0.59.5 ± 0.339.5 ± 1.0No inhibition5.9 ± 0.321.8 ± 0.46.4 ± 0.6
Beneckea natriegens9.1 ± 0.510.3 ± 0.214.1 ± 0.36.8 ± 0.37.3 ± 0.720.8 ± 1.86.2 ± 0.4
Brevibacterium linens6.7 ± 0.49.1 ± 0.521.7 ± 0.5No inhibition7.5 ± 0.312.7 ± 0.17.3 ± 0.6
Brocothrix thermosphacta7.4 ± 0.4No inhibition25.5 ± 1.0No inhibition6.1 ± 0.114.1 ± 0.27.4 ± 0.9
Citrobacter freundiiNo inhibitionNo inhibition17.7 ± 0.1No inhibition6.9 ± 0.19.1 ± 0.39.2 ± 0.2
Clostridium sporogenesNo inhibition16.6 ± 0.520.3 ± 0.78.9 ± 0.512.6 ± 0.29.7 ± 0.112.9 ± 0.5
Enterococcus faecalisNo inhibition11.3 ± 0.721.2 ± 0.4No inhibition21.6 ± 0.110.0 ± 0.112.9 ± 0.9
Enterobacter aerogenesNo inhibition12.0 ± 0.818.5 ± 0.8No inhibition6.2 ± 0.79.9 ± 0.16.4 ± 0.5
Erwinia carotovoraNo inhibition11.0 ± 0.415.5 ± 0.7No inhibition10.2 ± 0.110.0 ± 0.58.1 ± 0.2
Escherichia coli6.7 ± 0.313.5 ± 1.229.2 ± 0.25.9 ± 0.611.0 ± 0.213.3 ± 0.29.7 ± 0.7
Flavobacterium suaveolens7.0 ± 0.110.9 ± 0.426.0 ± 1.85.1 ± 0.56.6 ± 0.111.6 ± 0.67.0 ± 0.6
Klebsiella pneumoniaeNo inhibition11.7 ± 0.623.6 ± 0.17.1 ± 0.38.8 ± 0.210.9 ± 0.3No inhibition
Lactobacillus plantarumNo inhibitionNo inhibition18.7 ± 0.76.3 ± 0.77.9 ± 0.421.5 ± 0.66.2 ± 0.4
Leuconostoc cremorisNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibition
Micrococcus luteusNo inhibition11.1 ± 0.426.6 ± 0.5No inhibition6.7 ± 0.411.7 ± 0.76.1 ± 0.1
Moraxella sp.No inhibition12.3 ± 0.721.6 ± 0.1No inhibition6.5 ± 0.410.1 ± 0.66.1 ± 0.1
Proteus vulgarisNo inhibition11.1 ± 0.426.5 ± 1.66.2 ± 0.16.9 ± 0.18.3 ± 0.35.6 ± 0.3
Pseudomonas aeruginosaNo inhibition10.6 ± 2.026.0 ± 0.4No inhibition6.6 ± 0.115.5 ± 0.65.7 ± 0.3
Salmonella pullorumNo inhibition13.8 ± 0.827.1 ± 0.75.0 ± 0.512.0 ± 0.112.9 ± 0.16.3 ± 0.2
Serratia marcescens5.4 ± 0.68.0 ± 1.222.5 ± 0.9No inhibition6.0 ± 0.622.9 ± 0.85.7 ± 0.1
Staphylococcus aureus6.9 ± 0.211.3 ± 0.620.2 ± 0.5No inhibition4.9 ± 0.111.5 ± 0.55.2 ± 0.1
Yersinia enterocoliticaNo inhibition15.4 ± 0.222.4 ± 1.2No inhibition9.0 ± 0.611.6 ± 0.48.0 ± 0.2

Plant material

The volatile oils used in this study were isolated by hydrodistillation using essential oil distillation apparatus (‘Quick Fit’, British Pharmacopoeia, BDH, UK) The individual phytoconstituents were purchased either from Sigma (UK) or Fluka (UK) Chemicals.

Bacterial strains

Twenty-five bacterial strains were used to assess the antibacterial properties of the test samples, nine Gram-positive and 16 Gram-negative bacteria. Twenty-four out of 25 bacterial strains were maintained on Iso-Sensitest agar slopes [CM 471] (Oxoid, UK) at room temperature. Clostridium sporogenes was maintained in cooked meat broth under anaerobic conditions. All strains were subcultured every 2 weeks. The sources of the strains used are listed in Table 1.

Assessment of inhibition of bacterial growth

The measurement of growth inhibition was carried out in agreement with the method of Deans & Ritchie (1987) using Iso-Sensitest agar. Cells from cultures grown on Iso-Sensitest slopes were inoculated using a sterile loop into fresh Iso-Sensitest broth and incubated overnight at 25 °C (10 ml volume, 105 ml−1 final concentration). In the case of the Clostridium culture, a universal containing 20 ml of meat extract broth was boiled for 20 min and allowed to cool in order to create anaerobic conditions, and subsequently was incubated with a loopful of broth from the original inoculated culture. Next, 1 ml amounts of each culture were pipetted into separate sterile Petri dishes to which 20 ml amounts of molten Iso-Sensitest agar (45 °C) were added. Once set, wells of 4 mm diameter were made in the centre of each agar plate using a Pharmacia gel punch (Uppsala, Sweden), into which 15 μl test substance was added. The plates were then left undisturbed to allow diffusion of the sample into the agar, and incubated inverted in the dark at 25 °C for 48 h. Following this, zones of growth inhibition were measured using Vernier calipers.

Results

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Antibacterial activity of plant volatile oils

The antibacterial activities of the plant volatile oils presented in Table 1 are in general agreement with previously reported studies on the volatile oils of P. nigrum ( Deans & Ritchie 1987; Ouattara et al. 1997 ), S. aromaticum ( Deans et al. 1995 ; Cai & Wu 1996; Hao et al. 1998 ; Smith-Palmer et al. 1998 ), P. graveolens ( Pattnaik et al. 1996 ), M. fragrans, O. vulgare ( Kivanc & Akgül 1986) and T. vulgaris ( Kivanc & Akgül 1986; Smith-Palmer et al. 1998 ). All the bacterial strains demonstrated some degree of sensitivity to the plant volatile oils tested, although the growth of a number of bacteria were uninhibited by specific volatile oils. Zaika (1988) proposed that Gram-positive bacteria are more resistant then Gram-negative bacteria to the antibacterial properties of plant volatile oils which is in contrast to the hypothesis proposed by Deans that the susceptibility of bacteria to plant volatile oils and the Gram reaction appears to have little influence on growth inhibition ( Deans & Ritchie 1987; Deans et al. 1995 ). The volatile oils of O. vulgare ssp. hirtum, P. nigrum, S. aromaticum and M. fragrans did appear to be equally effective against both Gram-positive and Gram-negative microorganisms, in contrast to Zaika (1988), Hussein (1990) and Smith-Palmer et al. (1998) . However, P. graveolens and T. vulgaris volatile oils appeared preferentially more active with respect to Gram reaction, exerting greater inhibitory activity against Gram-positive organisms.

Table 1 summarizes the antibacterial activity of the volatile oils. From this, the oil with the widest spectrum of activity was found to be T. vulgaris, followed by O. vulgare ssp. hirtum, S. aromaticum, M. fragrans, P. nigrum, P. graveolens, in that order. Table 2 summarizes the antibacterial activity of the individual oil components. From this, the component with the widest spectrum of activity was found to be thymol followed by carvacrol, α-terpineol, terpinen-4-ol, eugenol, (±)-linalool, (–)-thujone, δ-3-carene, cis-hex-3-an-1-ol, geranyl acetate, (cis+ trans) citral, nerol, geraniol, menthone, β-pinene, R(+)-limonene, α-pinene, α-terpinene, borneol, (+)-sabinene, γ-terpinene, citronellal ∼ terpinolene, 1,8-cineole, bornyl acetate, carvacrol methyl ether, myrcene, β-caryophyllene, α-bisabolol, α-phellandrene, α-humulene, β-ocimene, aromadendrene, p-cymene, in that order.

Table 2b.  Zones of growth inhibition (mm) showing antibacterial activity for a number of selected plant volatile oil components; well diameter 4.0 mm
Bacterial strain891011121314
  1. Values for zone of growth inhibition are presented as mean ± SEM. 8, Geranyl acetate; 9, cis-hex-3-en-1-ol; 10, R(+)-limonene; 11, (±)-linalool; 12, menthone; 13, nerol; 14, α-pinene.

Acinetobacter calcoacetica10.3 ± 0.38.1 ± 0.1No inhibition9.3 ± 0.59.7 ± 2.311.4 ± 0.5No inhibition
Aeromonas hydrophila9.0 ± 0.48.5 ± 0.3No inhibition11.5 ± 0.97.0 ± 0.47.7 ± 0.1No inhibition
Alcaligenes faecalis10.5 ± 0.19.3 ± 0.6No inhibition12.1 ± 0.46.2 ± 0.57.1 ± 0.4No inhibition
Bacillus subtilis10.8 ± 0.26.4 ± 0.7No inhibition14.0 ± 0.87.1 ± 0.312.4 ± 0.2No inhibition
Beneckea natriegens10.8 ± 0.17.6 ± 0.5No inhibition11.4 ± 0.35.9 ± 0.411.3 ± 0.5No inhibition
Brevibacterium linens12.5 ± 0.88.1 ± 0.2No inhibition12.5 ± 0.7No inhibition11.7 ± 0.6No inhibition
Brocothrix thermosphacta9.2 ± 0.224.0 ± 0.6No inhibition8.1 ± 0.46.8 ± 0.49.0 ± 0.9No inhibition
Citrobacter freundii6.8 ± 0.89.7 ± 0.17.8 ± 0.127.5 ± 1.97.8 ± 0.67.8 ± 0.46.0 ± 0.3
Clostridium sporogenes20.4 ± 0.47.8 ± 0.210.3 ± 0.120.3 ± 0.410.7 0.3No inhibition5.7 ± 0.1
Enterococcus faecalis7.5 ± 0.68.9 ± 1.1No inhibition16.7 ± 1.1No inhibitionNo inhibition9.2 ± 0.1
Enterobacter aerogenes7.6 ± 0.26.5 ± 0.27.1 ± 0.29.7 ± 0.56.3 ± 0.17.2 ± 0.5No inhibition
Erwinia carotovora8.7 ± 1.29.3 ± 0.97.4 ± 0.112.3 ± 0.86.5 ± 0.57.7 ± 1.28.7 ± 0.7
Escherichia coli11.0 ± 0.212.0 ± 0.811.2 ± 0.313.8 ± 0.36.6 ± 0.27.6 ± 0.68.9 ± 0.5
Flavobacterium suaveolens11.0 ± 0.610.5 ± 0.210.6 ± 0.115.7 ± 2.45.8 ± 0.37.0 ± 0.46.5 ± 0.8
Klebsiella pneumoniae7.8 ± 0.410.7 ± 0.57.0 ± 0.112.6 ± 0.35.9 ± 0.4No inhibition8.1 ± 0.1
Lactobacillus plantarum12.9 ± 1.716.7 ± 2.9No inhibition25.3 ± 0.98.8 ± 0.719.1 ± 0.1No inhibition
Leuconostoc cremorisNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibitionNo inhibition
Micrococcus luteus8.0 ± 0.912.8 ± 0.8No inhibition13.4 ± 0.87.1 ± 0.37.4 ± 0.37.6 ± 0.8
Moraxella sp.9.0 ± 0.66.7 ± 0.27.9 ± 0.410.3 ± 0.96.9 ± 0.5No inhibition6.2 ± 0.4
Proteus vulgaris9.8 ± 0.18.2 ± 0.17.4 ± 0.512.2 ± 0.96.2 ± 0.1No inhibition7.5 ± 0.1
Pseudomonas aeruginosa6.5 ± 0.38.4 ± 0.3No inhibitionNo inhibitionNo inhibition13.6 ± 1.0No inhibition
Salmonella pullorum8.7 ± 0.412.0 ± 0.711.2 ± 0.67.5 ± 0.56.2 ± 0.6No inhibition7.9 ± 0.5
Serratia marcescens6.8 ± 0.112.4 ± 0.96.5 ± 0.18.8 ± 0.17.1 ± 0.48.5 ± 1.0No inhibition
Staphylococcus aureus6.6 ± 0.68.2 ± 0.3No inhibition9.0 ± 0.410.2 ± 1.09.4 ± 0.48.3 ± 0.1
Yersinia enterocolitica8.2 ± 1.011.5 ± 1.17.1 ± 0.29.5 ± 0.98.0 ± 0.27.1 ± 0.26.6 ± 0.6

Discussion

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The activity of the oils would be expected to relate to the respective composition of the plant volatile oils, the structural configuration of the constituent components of the volatile oils and their functional groups and possible synergistic interactions between components. A correlation of the antimicrobial activity of the compounds tested and their relative percentage composition in the plant volatile oils used in this study, with their chemical structure, functional groups and configuration, suggests a number of observations.

The components with phenolic structures, such as carvacrol, eugenol and thymol, were highly active against the test microorganisms. Members of this class are known to be either bactericidal or bacteriostatic agents, depending upon the concentration used ( Pelczar et al. 1988 ). These compounds were strongly active despite their relatively low capacity to dissolve in water, which is in agreement with published data ( Nadal et al. 1973 ; Suresh et al. 1992 ; Lattaoui & Tantaoui-Elaraki 1994; Mahmoud 1994; Meena & Sethi 1994; Shapiro et al. 1994 ; Belaiche et al. 1995 ; Jeongmok et al. 1995 ; Charai et al. 1996 ; Sivropoulou et al. 1996 ; Hili et al. 1997 ; Lis-Balchin & Deans 1997).

The importance of the hydroxyl group in the phenolic structure was confirmed in terms of activity when carvacrol was compared to its methyl ether. Furthermore, the relative position of the hydroxyl group exerted an influence upon the components effectiveness as seen in the difference in activity between carvacrol and thymol against Gram-negative and Gram-positive bacteria. Furthermore, the significance of the phenolic ring was demonstrated by the lack of activity of the monoterpene cyclic hydrocarbon p-cymene. The high activity of the phenolic components may be further explained in terms of the alkyl substitution into the phenol nucleus, which is known to enhance the antimicrobial activity of phenols ( Pelczar et al. 1988 ). The introduction of alkylation has been proposed to alter the distribution ratio between the aqueous and the nonaqueous phases (including bacterial phases) by reducing the surface tension or altering the species selectivity. Alkyl substituted phenolic compounds form phenoxyl radicals which interact with isomeric alkyl substituents ( Pauli & Knobloch 1987). This does not occur with etherified/ esterified isomeric molecules, possibly explaining their relative lack of activity.

The presence of an acetate moiety in the structure appeared to increase the activity of the parent compound. In the case of geraniol, the geranyl acetate demonstrated an increase in activity against the test microorganisms ( Table 2). Only Cl. sporogenes was found to be more resistant to the acetate. A similar tendency was identified in the case of borneol and bornyl acetate ( Table 2). Borneol was less active then the acetate except against Aeromonas hydrophila, Bacillus subtilis, Beneckea natriegens, Escherichia coli, Flavobacterium suaveolens and Serratia marcescens but only the acetate was capable of affecting the growth of the bacterium Micrococcus luteus.

Alcohols are known to possess bactericidal rather than bacteriostatic activity against vegetative cells. The alcohol terpenoids in this study did exhibit activity against the test microorganisms, potentially acting as either protein denaturing agents ( Pelczar et al. 1988 ), solvents or dehydrating agents.

Aldehydes, notably formaldehyde and glutaraldehyde, are known to possess powerful antimicrobial activity. It has been proposed that an aldehyde group conjugated to a carbon to carbon double bond is a highly electronegative arrangement, which may explain their activity ( Moleyar & Narasimham 1986), suggesting an increase in electronegativity increases the antibacterial activity ( Kurita et al. 1979 , 1981). Such electronegative compounds may interfere in biological processes involving electron transfer and react with vital nitrogen components, e.g. proteins and nucleic acids and therefore inhibit the growth of the microorganisms. The aldehydes cis + trans citral displayed moderate activity against the test microorganisms while citronellal was only active against B. subtilis, Cl. sporogenes, Fl. suaveolens, M. luteus and Pseudomonas aeruginosa ( Table 2).

A number of the components tested are ketones. The presence of an oxygen function in the framework increases the antimicrobial properties of terpenoids ( Naigre et al. 1996 ). From this study, and by using the contact method, the bacteriostatic and fungistatic action of terpenoids was increased when carbonylated. Menthone was shown to have modest activity, Cl. sporogenes and Staphphyloccus aureus being the most significantly affected ( Table 2).

An increase in activity dependent upon the type of alkyl substituent incorporated into a nonphenolic ring structure appeared to occur in this study. An alkenyl substituent (1-methylethenyl) resulted in increased antibacterial activity, as seen in limonene [1-methyl-4-(1-methylethenyl)-cyclohexene], compared to an alkyl (1-methylethyl) substituent as in p-cymene [1-methyl-4-(1-methylethyl)-benzene]. As shown in Table 2, the inclusion of a double bond increased the activity of limonene relative to p-cymene, which demonstrated no activity against the test bacteria. In addition, the susceptible organisms were principally Gram-negative, which suggests alkylation influences Gram reaction sensitivity of the bacteria. The importance of the antimicrobial activity of alkylated phenols in relation to phenol has been previously reported ( Pelczar et al. 1988 ). Their data suggest that an allylic side chain seems to enhance the inhibitory effects of a component and chiefly against Gram-negative organisms.

Furthermore, the stereochemistry had an influence on bioactivity. It was observed that α-isomers are inactive relative to β-isomers, e.g. α-pinene; cis-isomers are inactive contrary to trans-isomers, e.g. geraniol and nerol; compounds with methyl-isopropyl cyclohexane rings are the most active; or unsaturation of the cyclohexane ring further increases the antibacterial activity, e.g. terpinolene, terpineol and terpineolene ( Hinou et al. 1989 ).

Investigations into the effects of terpenoids upon isolated bacterial membranes suggest that their activity is a function of the lipophilic properties of the constituent terpenes ( Knobloch et al. 1986 ), the potency of their functional groups and their aqueous solubility ( Knobloch et al. 1988 ). Their site of action appeared to be at the phospholipid bilayer, caused by biochemical mechanisms catalysed by the phospholipid bilayers of the cell. These processes include the inhibition of electron transport, protein translocation, phosphorylation steps and other enzyme-dependent reactions ( Knobloch et al. 1986 ). Their activity in whole cells appears more complex ( Knobloch et al. 1988 ). Although a similar water solubility tendency is observed, specific statements on the action of single terpenoids in vivo have to be assessed singularly, taking into account not only the structure of the terpenoid, but also the chemical composition of the cell wall ( Knobloch et al. 1988 ). The plant extracts clearly demonstrate antibacterial properties, although the mechanistic processes are poorly understood. These activities suggest potential use as chemotherapeutic agents, food preserving agents and disinfectants.

Chemotherapeutic agents, used orally or systemically for the treatment of microbial infections of humans and animals, possess varying degrees of selective toxicity. Although the principle of selective toxicity is used in agriculture, pharmacology and diagnostic microbiology, its most dramatic application is the systemic chemotherapy of infectious disease. The tested plant products appear to be effective against a wide spectrum of microorganisms, both pathogenic and nonpathogenic. Administered orally, these compounds may be able to control a wide range of microbes but there is also the possibility that they may cause an imbalance in the gut microflora, allowing opportunistic pathogenic coliforms to become established in the gastrointestinal tract with resultant deleterious effects. Further studies on therapeutic applications of volatile oils should be undertaken to investigate these issues, especially when considering the substantial number of analytical studies carried out on these natural products.

The volatile oils and their component volatility and lack of solubility make these plant extracts less appealing for general disinfectant applications. However, a role as disinfectants of rooms has been reportedly investigated in a classical study ( Kellner & Kober 1954). Their volatility would be a distinct advantage in lowering microbial contamination in air and on difficult to reach surfaces. Although the minimum inhibitory concentrations for a selection of oils tested in a closed chamber were lower in the vapour phase ( Inouye et al. 1983 ), evidence suggests that such applications may have merit ( Taldykin 1979; Makarchuk et al. 1981 ).

As food preservatives, volatile oils may have their greatest potential use. Spices, which are used as integral ingredients in cuisine or added as flavouring agents to foods, are present in insufficient quantities for their antimicrobial properties to be significant. However, spices are often contaminated with bacterial and fungal spores due to their volatile oil content, often with antimicrobial activity, being enclosed within oil glands and not being released onto the surface of the spice matter. Volatile oils, which often contain the principal aromatic and flavouring components of herbs and spices, if added to foodstuffs, would cause no loss of organoleptic properties, would retard microbial contamination and therefore reduce the onset of spoilage. In addition, small quantities would be required for this effect. Furthermore, evidence suggests that these oils possess strong antioxidant activities ( Dorman 1999; Youdim et al. 1999 ), which are favourable properties to combat free radical-mediated organoleptic deterioration.

Acknowledgements

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

H.J.D.D. gratefully acknowledges financial support through a MAFF Postgraduate Studentship. S.A.C. received financial support from the Scottish Office Agriculture Environment and Fisheries Department.

References

  1. Top of page
  2. Abstract
  3. MATERIALS and METHODS
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
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