3Division of Bacteriology and Mycology, ICAR-Indian Veterinary Research Institute, India
Received date: 15/10/2015 Accepted date: 19/11/2015 Published date: 25/11/2015
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Gum Acacia, Gum Chironji, Gum Karaya, Gum Balanites aegyptiaca, Gum Prosopis juliflora, Salmonella enterica, Escherichia coli, Klebsiella pneumonia, Brucella abortus, Bordetella bronchiseptica, Pasterurella multocida.
Edible gums known by different names viz., Gond, Goond, Goond katira, Dinka, Gaund, Gondh are dried sap (exudates) of thorny trees and shrubs. Gums from trees and shrubs of Fabaceae family (Acacia, Sterculia, Astragalus, Balanites, Buchanania, Anogeissus species) are edible . Edible gums are water soluble producing viscous gel like solution. They are used in food industry as thickening, gelling, emulsifying and stabilizing agents . Due to their adhesives, binding, clarifying, encapsulating, flocculating, swelling and foam stabilizing properties gums have multiple applications in multiple industries [1,2]. Gums are ionic polymers (polyelectrolytes) of acidic glycopeptides containing several biologically active compounds and mineral salts . Commonly available edible gums come from Acacia spp. trees and are known as gum acacia, chaar gund, char goond, or meska . It is mostly obtained from trees of two Acacia species viz., A. senegal and A. seyal. Other edible gums available in market are; gum ghatti (from Anogeissus trees), gum tragacanth (from Astragalus shrubs, gond katira), karaya gum (from Sterculia trees), gum babul (from Acacia nilotica), gum chironji (from Buchanania lanzan trees), and gums from Balanites aegyptiaca and Prosopis juliflora are also edible [1,4].
Antimicrobial activity of gums is always being a controversial issue due to several contrasting reports . Alcoholic extract and aqueous extracts of gum acacia are reported to inhibit growth of Staphylococcus aureus, S. epidermidis, Streptococcus pneumoniae, Pseudomonas aeroginosa, Proteus merabilis, Acinetobacter, Enterobacter, Klebsiella pneumoniae, Serratia spp., E. coli, Salmonella typhi, Cerospora pongamae, Candida albicans and Aspergillus niger [5,6]. Though gum chironji is reported beneficial in intercostals’ pain and diarrhoea, reports are scant on its antimicrobial activity . Antibacterial and antimycotic activity has been reported in alcoholic extracts of Balanites aegyptiaca leaves, bark and fruit mesocarps [7-11] inhibiting of metallo- B-lactmase producing E. coli, Klebsiella spp. and Citrobacter spp.  Though hot and cold aqueous extracts of Prosopis juliflora (Vilayati kikar) leaves inhibited Bacillus subtilis, E. coli, Enterobacter faecalis, K. pneumoniae, P. aeruginosa, S. aureus, S. epidermidis, S. pyogenes, S. typhi and S. typhimurium , its gum has rarely been reported to have antimicrobial activity. Gum Karaya from Sterculia urens tree has been used as an adhesive for dental fixtures and osteomyo- equipment, and as a base for salicylic acid patches [13,14], and is known to reduce bacterial adhesion by 98% when applied as protective coating to dentures  but its antibacterial activity is not reported yet. Antimicrobial activity of oleoresins (inedible gums) also similar to edible gums in texture is often reported , information on antimicrobial activity of edible gums is scanty and mostly based on tests using reference laboratory strains . The scant information on antimicrobial activity that too not on clinically important pathogens, limits the understanding of utility of edible gums as natural antimicrobials. This study aimed at determining antimicrobial activity of ethanolic extracts and aqueous solutions of commonly available edible gums in India on clinically important bacteria.
A total of 31 samples (duly authenticated and catalogued at ICAR-Indian Institute of Natural Resins and Gums (IINRG), Namkum, Ranchi) of Acacia nilotica (Babul) gum (17), Buchanania lanzan (Chironji) gum (7), Sterculia (Karaya) gum (5), Balanites aegyptiaca gum (1) and Prosopis juliflora gum (1) were collected/procured from fresh collections (gathered from different parts of the country) available at IINRG, Ranchi (Table 1). All the gum samples after manual cleaning and sorting were converted into fine powder and passed through 0.4 mm mesh sieve and packed in air tight containers for further analysis.
|Gum type||Place of collection||Serial numbers of samples collected|
|Gum Acacia||Sitapur, Uttar Pradesh||1|
|Balaghat, Madhya Pradesh||8|
|Jabalpur, Madhya Pradesh||9|
|Jodhpur, Rajasthan||13, 14|
|Gum Chironji||Bilaspur, ChattisGarh||16|
|Simdega, Ranchi, Jharkhand||17|
|ICAR-IINRG Farm, Ranchi, Jharkhand||18a|
|Dindori , Madhya Pradesh||18b|
|Umaria, Madhya Pradesh||19, 20|
|Sitapur, Uttar Pradesh||21|
|Gum Balanitesaegyptiaca||Jodhpur, Rajasthan||22|
|Gum Prosopisjuliflora,||Jodhpur, Rajasthan||23|
|Gum Karaya||Girijan Cooperative Corporation Ltd., Visakhaptnam, Andhra Pradesh||26-30|
Table 1: Source wise distribution of gum samples tested for antimicrobial activity in the study.
Ethanolic extracts of gums
To make Ethanolic extract (EE), 250 g of powdered gum was mixed in a 1 L conical flask with 500 ml of 99.9% pure ethanol (SD Fine Chem Ltd, Mumbai). Flasks were kept overnight at 25°C, over a rotary (30 rpm) platform. Thereafter, all the contents of flasks were filtered through glass wool filter to recover the filtrate. Filtrate was transferred to sterile shallow bowls and ethanol was allowed to evaporate at 50°C in an oven with exhaust overnight. Next morning all the dried material available in bowls was collected, weighed and kept in airtight screw capped vials at 4°C till tested within a month. To test the antimicrobial activity, discs of EE were prepared by dissolving 300 mg of EE in to 3 ml of ethanol and then adsorbing the solution on to 6 mm sterile discs (20 μl each), discs were allowed to dry at room temperature in desiccators and then stored in dry and sterile vials at 4°C till used within a month.
Aqueous extract (solution) of gums
To make gum solution in water (4% w/v), 2 gm of gum was added to 50 ml of sterile water in a sterilized conical flask and solubilized through keeping flasks in warm (50°C) shaking (50 rpm) water bath for 30 min. All gum solutions were tested on the same day for antimicrobial activity using agar well diffusion assay .
Four reference strains, sensitive to all commonly used antimicrobials (Streptococcus milleri SM-22; Bacillus mycoides B29- 19-1; E. coli E-382; Salmonella abortusequi E-155), available in Epidemiology Laboratory of Indian Veterinary Research Institute, were revived from glycerol stocks and tested for purity and identity and maintained on nutrient agar slants without further subculturing throughout the study . Besides, 97 isolates from clinical and environmental sources (Table 2) maintained in glycerol stocks and isolated during last three months at Clinical Epidemiology Laboratory of the Division, belonging to 24 species of 6 genera of Gram positive (34) bacteria (GPB) and to 27 species of 19 genera of Gram negative (63) bacteria (GNB) were included in the study to screen the antibacterial activity of EE and aqueous solutions of gums.
|Bacteria tested (no. of strains tested)||Number of strains sensitive||MIC of EE in µg/ml|
|Bacillus cereus (1)||1||1||0||1||80||80||>2560|
|Escherichia coli (14)||1||0||0||12||1280->2560||>2560||>2560|
|Proteus mirabilis (2)||1||1||0||1||320||640||>2560|
|Proteus vulgaris (1)||0||0||0||2||>2560||>2560||>2560|
|Pseudomonas aeruginosa (2)||0||0||0||2||>2560||>2560||>2560|
|Staphylococcus aureus ssp.(1)||0||0||0||1||>2560||>2560||>2560|
|Strept. equi ssp. equisimilis(1)||1||1||1||1||80||80||160|
|Total (101) sensitive strains (%)||50 (49.5)||49 (48.5)||9 (8.9)||88 (87.1)||80 to >2560||80 to >2560||160 to >2560|
|EE, ethanolic extract (EE); GA9, 2 mg discs of gum Acacia EE sample number 9; GA4, 2 mg discs of gum Acacia EE sample number 4; GC19, 2 mg discs of gum Chronji EE sample number 19, Cip10, ciprofloxacin discs (10 µg).|
Table 2: Antimicrobial activity and minimum inhibitory concentration of alcoholic extracts of gum Acacia (Acacia nilotica) and gum Chironji (Buchanania lanzan) against bacterial strains of public health concern.
Testing antimicrobial activity
Disc diffusion assay using EE discs was performed as described earlier [16,18] for different bacteria in triplicate on Mueller Hinton agar (MHA, BD BBL and Difco) plates, swab inoculated with overnight broth culture (0.1 OD590). All strains were tested for sensitivity on MHA but Moraxella, Streptococcus, Brucella, Bordetella and Pasteurella (slow growing, fastidious) strains were tested on Brain heart infusion (BHI) agar (BD BBL and Difco) instead of MHA . All strains were tested at 37°C aerobically except Brucella, which were incubated at 37°C in 5% CO2 enriched environment. Ciprofloxacin (10 μg) discs (BD BBL and Difco), and blank discs (first soaked in ethanol and dried in similar way as for EE discs) were used as standard antimicrobial and negative control discs, respectively.
Determination of MIC of EE of gum samples
It was performed using agar well diffusion assay using suitable agar plates (MHA/ BHIA) having 9 wells bored in each plate . Plates were swab inoculated with test strain as described for disc diffusion assay and then 50 μl of EE solution in dimethyl sulphoxide (DMSO, Merck India) was poured into each well. In first well DMSO without EE was used as negative control and in 2nd to 9th well 50 μl DMSO containing 20, 40, 80, 160, 320, 640, 1280 μg and 2560 μg of gum EE, respectively was poured in. Plates were incubated without inverting for 3 hr. so that contents of the well get diffused into the surrounding medium and then incubated as for disc diffusion assay. After 24 hr. of incubation plates were observed for inhibition of growth around the wells. The inhibition of growth around the wells with minimum EE was considered MIC of the EE of the gum. If no inhibition of bacteria was observed in a plate then MIC was recorded as >2560 μg/ml.
To determine correlation between diameter of zone of inhibition (in mm) of bacteria around EE discs and MIC, correlation coefficient was calculated using MS Office Excel-7. To estimate association between sensitivity of bacteria to EE of gums and species of bacteria, χ2 test was performed in MS Office Excel-2007. The statistical comparison was done for only those genera of bacteria where number (n) of strains tested of the genus was ≥ 10.
None of the 101 strains including 4 reference (Streptococcus milleri SM-22; Bacillus mycoides B29-19-1; E. coli E-382; Salmonella abortusequi E-155) strains was inhibited by aqueous solution of any of the 31 gum samples. Discs of EE of only three gum samples (no. 4 and 9 of gum acacia and no. 19 of gum chironji) induced variable zone of growth inhibition (Table 2). All the three EEs inhibiting growth of bacteria failed to inhibit growth of both of the two reference GNB strains while both of the reference GPB strains were inhibited by all the three EEs having antimicrobial activity. Of the 101 strains, 50 strains (49.5%) were sensitive to EEA-9, 49 (48.5%) to EEA-4 and 9 (8.9%) to EEC-19. There was near to perfect (r=0.87) correlation between inhibition zones caused by EEA-4 and EEA-9 discs for different bacteria.
Including 4 reference strains, a total of 88 (87.1%) strains were sensitive to ciprofloxacin disks. Of the 36 GPBs tested, 26 were sensitive to EEA-4 and EEA-9, 4 to EEC-19 and 28 to ciprofloxacin disks. Of the 65 GNB strains tested, 60, 24, 23 and 5 were sensitive to ciprofloxacin, EEA-9, EEA-4, and EEC-19 disks, respectively.
Sensitivity to EEA was more common among GPBs (p=0.0007) than GNBs but no significant difference was evident for EEC (p=0.56). Ciprofloxacin resistance was more (p=0.04) common in GPB than in GNB strains.
Minimum inhibitory concentration (MIC) for EEA from gum acacia sample no. 4 and 9 for sensitive strains ranged between 80 μg to 1280 μg/ml for GNBs and 80 μg to 640 μg for GPBs (Table 2). For sensitive strains MIC was minimum (80 μg/ml) for Aeromonas bestiarum, Moraxella osloensis, P. canis and Streptococcus equi and was maximum (2560 μg/ml) for Actinobacillus actinomycetencomitans strains. MIC of EEC-19 was the lowest (160 μg) for Strept. equi and maximum (2560 μg/ml) for Acinetobacter baumanii strains, and ranged between 640 μg to 1280 μg/ml for other sensitive strains. For all the strains resistant to EEA and EEC MIC was >2.56 mg/ml (Table 2).
Natural edible gums though reported to be antimicrobial in some reports [5,6], are generally considered devoid of antimicrobial activity . In presented study three of the 31 gum samples had detectable antibacterial activity in concurrence of earlier observations . Variations reported in composition of different gum samples [1,3,4] might be responsible for antimicrobial activity in EE of three gum samples and not in other gum samples. Antibacterial activity only in alcoholic extracts but not in aqueous solution of gums was in agreement to earlier studies [5,6,11,19-22]. The study also revealed that EE of acacia gum sample no. 9 (from Jabalpur) was a little better in inhibiting growth of 50 strains of bacteria than EE of acacia gum sample no. 4 from Rohtak (inhibiting only 49), but the difference was statistically insignificant and MIC studies also indicated the similar MIC of both the EEAs for different bacteria. The difference in antimicrobial activity of gum acacia samples cannot be explained on the basis of source (geographical region) as in the same region samples without antimicrobial activity were available. The difference might be attributed to individual plant source but it needs further studies for verification and elucidation of exact reason of variation.
Wide spectrum of activity of EE of gum acacia against 50 strains of bacteria belonging to 35 species of GPBs and GNBs both indicated presence of a promising antimicrobial compound in gum acacia. In earlier studies gum acacia extracts are reported to inhibit growth of 12 reference strains of S. aureus, S. epidermidis, S. pneumoniae, P. aeroginosa, Proteus merabilis, Acinetobacter, Enterobacter, Klebsiella pneumoniae, Serratia spp., E. coli, Salmonella typhi, Candida albicans  and reference strains of Bacillus cereus, E. coli, Aspergillus niger and Cercospora pongamiae  indicating wide spectrum of its antimicrobial activity. However, this study seems to be first to report the antimicrobial activity of gum acacia extracts on clinically important bacteria. Moreover, from earlier studies [6,23] variation in sensitivity of different strains of the same bacteria was not apparent, this study elucidated that only small fraction of E. coli, Proteus, Aeromonas, Staphylococcus and Streptococcus strains was susceptible to antimicrobial action of gum acacia. The present study suggested that conclusion based on studies using only a few reference strains may not be always practically useful. Further, detection of sensitivity in a few strains of a bacteria or resistance in a few strains of other bacteria indicated that there might be some mechanism for emergence of resistance against herbal antimicrobials as proposed earlier [16,24-26].
Antimicrobial activity in gum chironji or its extract has not been reported earlier [1,19]. In the present study one of the 7 gum samples had some antimicrobial potential that too against a few strains of Aeromonas veronii, B. mycoides, Brucella abortus and S. milleri indicating its weak antimicrobial potential.
Lack of detection of any antimicrobial activity in gum samples of Sterculia (Karaya), Balanites aegyptiaca and Prosopis juliflora is in concurrence of the earlier studies [1,11,12,27-29]. It might be either due to the absence of antimicrobial potential in those edible gums or due to less number of samples analyzed.
All the three gum samples having antimicrobial activity in their alcoholic extract were from three different regions (Table 1) but from the same or nearby regions other samples of the same gum were negative for antimicrobial potential. For the observation no reason could be assigned on the basis of present study and it needs to be explored further.
Though edible gums may not be considered useful antimicrobials for therapeutic use , utility of gums in green synthesis of nanoparticles, nano-fibers for antimicrobial scaffolds useful in dressing and nano-capsules for drug delivery or membranes of packaging is emerging fast [27,30-34]. Moreover, further studies may lead to identification of active antimicrobial component of gum which might be a potential one.
Although, antimicrobial activity in acacia gum has wide spectrum, was detected in only two of the 17 samples. Similarly, only one of the 7 gum chironji samples possessed a little antimicrobial activity. To elucidate the reasons behind antimicrobial potential only in a few gum samples and for identification of active antimicrobial component in edible gum samples need further studies. Moreover, identification of the active antimicrobial substance in alcoholic extracts of gum acacia and gum chironji might be important area of further research.
Authors are thankful to Director, Joint Director (R) and Joint Director (A) of ICAR-Indian Veterinary Research Institute, Izatnagar for permitting for the study and grant of fellowship to Sakshi Dubey, M. Senthil Murugan, Polem Lenin Singh, Prasanna Vadhana A and Monika Bhardwaj and Director ICAR-IINGR, Ranchi for permission to work for preparation and procurement of gum samples. Authors also thank Mr. HC Joshi and Mr. Laikurahman for consistent laboratory maintenance activities.