Clostridium difficile infection (CDI) is a leading cause of antibiotic-associated diarrhea in health care facilities in developed countries. Extended hospital stays and recurrences severely increase the cost of treatments and the high mortality rate that is observed among the elderly. Community-associated CDI cases that occur without any recent contact with the hospital environment are increasing. Studies have reported the isolation of virulent C. difficile strains from water, soil, meat, vegetables, pets, livestock animals, and animal manure. The objective of this study was to isolate and characterize C. difficile strains from animal manure and commercially available compost products. Our results demonstrate that not only unprocessed animal manure but also finished composts made of different feedstocks can serve as a reservoir for C. difficile as well. Most importantly, our study revealed that properly processed compost is a potential source of toxigenic and clindamycin-resistant C. difficile isolates.
KEYWORDS: C. difficile, animal manure, compost, toxigenicity, ribotyping, antimicrobial resistance
ABSTRACT
The well-known nosocomial pathogen Clostridium difficile has recently been recognized as a community-associated pathogen. As livestock animals carry and shed C. difficile in their feces, animal manure-based composts may play an important role in disseminating toxigenic C. difficile strains into the agricultural environment. The present study surveyed C. difficile contamination of commercially available composts and animal manure. Presumptive C. difficile isolates were confirmed by testing for the tpi housekeeping gene in addition to Gram staining. The confirmed C. difficile isolates were further tested for toxigenicity, PCR ribotype, and susceptibilities to selected antibiotics. C. difficile was found in 51 out of 142 samples (36%). A total of 58 C. difficile strains were isolated from those 51 positive compost/manure samples. The presence of C. difficile in compost did not significantly correlate (P > 0.05) with the physical and most microbiological parameters, including the presence of fecal coliforms. The majority of C. difficile isolates were toxigenic, with 63.8% positive for the toxin A gene (tcdA) and 67.2% positive for the toxin B gene (tcdB). Only 3 isolates (5.17%) were positive for the binary toxins. There were 38 different PCR ribotypes among the 58 C. difficile isolates, and ribotype 106 was the most prevalent, followed by ribotypes 020 and 412. All C. difficile isolates were susceptible to the selected antibiotics, but >50% of the isolates had reduced susceptibility to clindamycin by the agar dilution method. This study indicates that compost may be a reservoir of toxigenic C. difficile strains.
IMPORTANCE Clostridium difficile infection (CDI) is a leading cause of antibiotic-associated diarrhea in health care facilities in developed countries. Extended hospital stays and recurrences severely increase the cost of treatments and the high mortality rate that is observed among the elderly. Community-associated CDI cases that occur without any recent contact with the hospital environment are increasing. Studies have reported the isolation of virulent C. difficile strains from water, soil, meat, vegetables, pets, livestock animals, and animal manure. The objective of this study was to isolate and characterize C. difficile strains from animal manure and commercially available compost products. Our results demonstrate that not only unprocessed animal manure but also finished composts made of different feedstocks can serve as a reservoir for C. difficile as well. Most importantly, our study revealed that properly processed compost is a potential source of toxigenic and clindamycin-resistant C. difficile isolates.
INTRODUCTION
The annual animal manure production in the United States is estimated to be 1.37 billion tons, including cattle, hog, chicken, and turkey manure (1). Due to the high nitrogen and phosphorus contents, animal manure and composted manure are used as crop fertilizers in agricultural production and to improve the soil structure (2). Animal manure is a well-known source of foodborne pathogens, such as Salmonella spp., Escherichia coli O157:H7, Campylobacter spp., Listeria monocytogenes, and parasites, such as Cryptosporidium parvum (2). Composting is an effective way to eliminate the majority of aforementioned human pathogens in animal manure before their use as fertilizers. During the thermophilic phase of composting, high temperatures generated by microbial activities can reach ca. 55 to 70°C inside the composting piles/heaps and are typically held for a few days to several weeks. The elevated temperatures have been proven to inactivate Gram-negative pathogens, such as E. coli O157:H7 and Salmonella spp., inside the static heaps of dairy manure within 3 weeks of composting (3). However, limited studies have assessed the fate of endospore-forming Gram positives, such as Clostridium difficile, in composting (4).
C. difficile species cause enteric diseases in humans and animals. Clostridium difficile infection (CDI) is mainly associated with exposure to antibiotics, which decreases the competition of gut microflora and allows endospore-forming C. difficile bacteria to proliferate. The symptoms can vary from mild diarrhea to life-threatening toxic megacolon or pseudomembranous colitis in humans (5). Infected animals/humans shed the vegetative cells and endospores of pathogenic clostridia with their fecal matter (6). The abundance of C. difficile in livestock animals was surveyed across 35 states in the United States, and it was reported that 16, 6.3, and 2.4% of fecal samples were positive for C. difficile in swine, beef cattle, and dairy cattle, respectively (6).
Although the sources of contamination have not been confirmed, evidence for C. difficile contamination in vegetables and ready-to-eat vegetable salad ingredients has been reported in Scotland, France, Canada, and the United States (7–10). All studies reported an abundance of <10% in those vegetables, but all isolates were human-pathogenic C. difficile strains. A few studies have reported that composting temperatures reduced the level of C. difficile endospores in biosolids and swine manure (4, 11). If a high initial load of endospores is present in unprocessed animal manure, it is possible for a fraction of those endospores to survive in finished compost due to the inherent heat resistance. The importance of evaluating the prevalence of C. difficile in manure, composts, and biosolids used for agricultural and recreational purposes was emphasized by Moono et al. (12), who detected a high prevalence of toxigenic C. difficile strains in public space lawns. Similarly, when contaminated compost is subsequently used to fertilize crops, it may contaminate the edible parts of fresh produce by C. difficile via wind, insects, and human activities. The ingestion of pathogenic clostridial endospores of toxigenic and antimicrobial-resistant strains may result in symptoms of CDI.
The objective of this study was to determine the contamination of animal manure/composts by C. difficile from the samples collected from multiple states across the United States. Further characterization of isolates was performed to broaden the current knowledge of potential sources of C. difficile as a community-associated pathogen.
RESULTS
Physical and microbiological parameter analysis.
For the convenience of reporting, finished compost samples were categorized into 6 types: (i) cow manure-based compost, (ii) poultry litter-based compost, (iii) plant-based compost (plant, raw, and cooked food waste), (iv) mixed manure-based compost (mixed, sheep, horse, and goat manure-based compost), (v) biosolid-based compost (both biosolid and municipal solid waste-based compost), and (vi) other composts (mushroom, peat moss, and fish emulsion based). There was another category for unprocessed manure (Table 1). The moisture content (MC) of all samples ranged approximately from 7.74 to 97.98%, with an overall average of 42.56%. The highest MC (97.98%) was for a composted tea product (spray), the only liquid sample analyzed in this study. Unprocessed manure had the highest average MC (56.20%), while the biosolid-based compost category had the lowest average MC (34.54%). The pH values of the majority of the samples were in the range of 3.88 to 9.73, with an average of pH 7.87. Cow manure and mixed-manure samples had pH >8.00. The average water activity (aw) for all compost/manure samples was 0.967, and the average aw for each category was >0.900.
TABLE 1.
Physical and microbiological property analysis results from compost/manure samples
| Source of compost/manure | n | MC (avg ± SD) (%) | pH (avg ± SD) | Water activity (avg ± SD) | Counts (median/avg) (log CFU/g) |
|||||
|---|---|---|---|---|---|---|---|---|---|---|
| Mesophiles | Mesoactinomycetes | Thermophiles | Thermoactinomycetes | Yeast and molds | Fecal coliformsa | |||||
| Cow manure based | 20 | 44.14 ± 21.66 | 8.27 ± 0.74 | 0.977 ± 0.450 | 7.91/7.86 | 7.64/7.40 | 6.81/6.70 | 6.32/6.24 | 4.68/4.28 | 2.83/3.26/40.0 |
| Poultry litter based | 20 | 40.79 ± 26.30 | 7.42 ± 3.42 | 0.929 ± 0.421 | 8.14/8.13 | 6.91/6.80 | 6.81/6.61 | 6.37/6.24 | 4.79/4.88 | 2.52/2.82/17.9 |
| Plant based | 25 | 40.91 ± 16.90 | 7.96 ± 0.88 | 0.981 ± 0.038 | 7.99/8.03 | 7.60/7.39 | 7.32/7.32 | 6.75/6.77 | 4.73/4.79 | 2.14/2.32/12.0 |
| Mix manure based | 26 | 43.91 ± 13.91 | 8.20 ± 0.74 | 0.985 ± 0.020 | 8.06/7.76 | 7.38/7.45 | 7.17/7.07 | 6.58/6.51 | 5.25/4.85 | 2.68/3.04/34.6 |
| Biosolid based | 22 | 34.54 ± 11.05 | 7.637 ± 0.97 | 0.940 ± 0.146 | 8.12/8.39 | 7.64/7.37 | 7.65/7.55 | 6.60/6.67 | 4.59/4.52 | 1.46/1.46/4.5 |
| Other composts | 8 | 52.18 ± 16.54 | 7.14 ± 2.19 | 0.988 ± 0.027 | 8.08/8.06 | 7.41/7.26 | 6.83/7.08 | 6.50/6.37 | 5.71/5.46 | 1.78/1.78/12.5 |
| Unprocessed manure | 21 | 56.20 ± 31.94 | 7.87 ± 3.70 | 0.964 ± 0.495 | 8.86/8.63 | 8.24/8.07 | 6.06/6.29 | 5.54/5.53 | 5.94/4.88 | 5.86/2.58/47.6 |
Third number is percent positive results among samples tested in each category.
Results of the microbiological analysis of manure/compost samples are summarized in Table 1. Thermophilic bacterial counts in samples ranged from approximately 2.82 to 9.47 log CFU/g. The lowest average (6.29 log CFU/g) for thermophilic bacteria was observed for unprocessed manure, while the highest average (7.55 log CFU/g) was observed for biosolid waste-based compost. Mesophilic bacterial counts for all samples varied from 5.21 to 10.09 log CFU/g. Average mesophilic counts of >8.00 log CFU/g were observed for all categories, except for cow and mixed-manure-based compost categories. Thermoactinomycete counts in analyzed samples ranged from approximately 3.26 to 8.21 log CFU/g. The lowest average (5.54 log CFU/g) for thermoactinomycetes was seen in unprocessed manure, while each finished compost category had >6 log CFU/g as the average. Mesoactinomycete counts for all samples varied from 5.00 to 9.40 log CFU/g. The average mesoactinomycete counts were >7.00 log CFU/g for each category, except for poultry litter-based composts. Yeast/mold counts ranged from 4.5 to 6.00 log CFU/g for samples analyzed in all of the categories. Fecal coliforms were detected in 23.14% (28/121) of the finished compost samples, with the highest proportion being from cow manure-based composts and the lowest being for biosolid waste-based composts. The fecal coliform level was below the detection limit (0.7 log CFU/g) for more than 70% (36/51) of the C. difficile-positive compost/manure samples.
Isolation of C. difficile from manure/compost.
C. difficile was isolated from 51 (47 from finished compost and 4 manure samples) out of 142 samples (36%) of manure/compost (Table 2). All categories of finished compost samples were positive for C. difficile. The highest positive rate for C. difficile was observed in biosolid waste-based compost (59.09%), and the lowest positive rate was in poultry litter-based compost (20.00%). The positive rate for unprocessed manure samples was 19.05%. Approximately 15% of C. difficile-positive finished compost samples (7/47) were positive for fecal coliforms as well, and only one (mixed-manure-based) of those 7 samples reported more than 3 log CFU/g compost for fecal coliforms. The statistical analysis showed that there were no significant (P > 0.05) correlations between the presence of C. difficile and the presence of fecal coliforms. Moisture content (P = 0.037), thermophilic count (P = 0.035), and mesoactinomycete count (P = 0.011) had significant effects on the presence of C. difficile in composts using the complete model for manure/compost samples. However, in the stepwise approach, not all factors significantly contributed to the presence of C. difficile in finished compost or unprocessed manure used in this study (P > 0.05). Only the mesoactinomycete count (P = 0.018) had a significant effect on the presence of C. difficile in finished compost, while the moisture content (P = 0.000), thermoactinomycete (P = 0.001), and mesoactinomycete (P = 0.004) counts were significant for the presence of C. difficile in unprocessed manure.
TABLE 2.
Summary of the distribution of C. difficile ribotypes among different feedstock of compost/manure samples
| Manure/compost type | No. of samples tested | No. (%) of C. difficile-positive samplesa | C. difficile ribotypes |
|---|---|---|---|
| Cow manure based | 20 | 8 (40.00) | 001, 012, 033, 075, 106, 596, 705, PR 13474 |
| Poultry litter based | 20 | 4 (20.00) | 009, 039/2, 106, PR 13476 |
| Plant based | 25 | 7 (28.00) | 014/0, 034, 039/2, 449, 106, PR11498, PR11692 |
| Mixed manure based | 26 | 12 (36.36) | 005, 009, 010, 020, 039/2, 081, 251, 412, 652 |
| Biosolid based | 22 | 13 (59.09) | 009, 010, 017, 020, 039/02, 078, 203, 235, 251, 404, 412, 106, 684, AI 8/0,AI 82/1, AI/83, PR 13475 |
| Other composts | 8 | 3 (37.5) | 449, 593, PR11665 |
| Unprocessed manure | 21 | 4 (19.05) | 115, 412, 446, 705 |
| Total | 142 | 51 (35.92) |
Percent positive among samples tested in each category.
Detection of toxin genes.
PCR was used to distinguish the toxin types A+B+, A−B+, and A−B− of C. difficile. The majority of the isolates were toxigenic, with 62.07 and 65.51% positivity rates for genes for toxins A (tcdA) and B (tcdB), respectively. From 2 isolates which were positive for only tcdB, one isolate had the deleted fragment of tcdA (110 bp), while the other isolate had neither tcdA nor deleted tcdA. There were 3 isolates positive for binary toxins. One of those isolates was negative for both tcdA and deleted tcdA, and other two isolates were positive for both toxins A and B.
Ribotyping.
Among the 58 C. difficile isolates, 38 different ribotypes were identified by sequence-based PCR-ribotyping (Tables 2 and 3). The fragment size of each isolate varied from 231 to 546 bp. The best match of the Webribo database was accepted when the band pattern, band sizes, and the genetic analyzer matched the experimental conditions. Among our isolates, the most common toxigenic (A+B+) ribotype was 106, and it was isolated from all categories of compost, except for the mixed compost and other compost types. Ribotypes 020 and 412 were the next most prevalent toxigenic ribotypes. The most prevalent nontoxigenic (A−B−) ribotype was 009. There were 8 ribotypes (8/38) that could not be assigned to an internationally recognized ribotype. The same ribotype was isolated from multiple compost samples made from different feedstocks, and some compost samples had more than one ribotype. From the 3 isolates positive for binary toxins, one isolate was identified as ribotype 078, which was the isolate from biosolid waste-based compost without either tcdA or deleted tcdA. The other two positive isolates for binary toxins were identified as ribotype 251 (A+B+CDT+).
TABLE 3.
Analysis of ribotypes and toxigenic genes of C. difficile isolated from compost/manure samples
| Sample IDa | Presence of toxin gene result |
Ribotypeb | |||
|---|---|---|---|---|---|
| tcdA | tcdB | cdtA | cdtB | ||
| ATCC 43593 | − | − | − | − | PR11586 (060) |
| ATCC BAA-2155 | + | + | + | + | PR11587 (251) |
| CPS 5 | + | + | − | − | PR11676 (012) |
| CPS 7 | + | + | − | − | PR11677 (652) |
| CPS 10 | + | + | − | − | PR 13477 (005) |
| CPS 14 | − | − | − | − | PR11678 (075) |
| CPS 15 | + | + | − | − | PR11567 (705) |
| CPS 16 | + | + | − | − | PR11493 (020) |
| CPS 17 | − | − | − | − | PR11660 (033) |
| CPS 22 | − | − | − | − | PR11679 (034) |
| CPS 23 | + | + | − | − | PR11589 (106) |
| CD 6 | + | + | − | − | PR11653 (446) |
| CD 13 | + | + | − | − | PR11688 (115) |
| CD 14 | − | − | − | − | PR11528 (009) |
| CD 22 | + | + | − | − | PR11689 (412) |
| CD 24 | + | + | − | − | PR11588 |
| CD 25 | − | − | − | − | PR13473 (596) |
| CD 28 | + | + | − | − | PR11567 (705) |
| CD 37 | − | − | − | − | PR13476 |
| CD 38 | + | + | − | − | PR11589 (106) |
| CD 41 | + | + | − | − | PR11657 (081) |
| CD 47 | + | + | − | − | PR11589 (106) |
| CD 49 | − | − | − | − | PR11656 (039/2) |
| CD 50 | − | − | − | − | PR13475 |
| CD 60 (I) | NDc | + | + | + | PR11562 (078) |
| CD 60 (II) | + | + | − | − | PR11681 (404) |
| CD 60 (III) | + | + | − | − | PR11682 (684) |
| CD 61 | − | − | − | − | PR11656 (039/2) |
| CD 70 | + | + | − | − | PR11589 (106) |
| CD 74 | + | + | − | − | PR11690 (002/2) |
| CD 75 | + | + | − | − | PR11691 (014/0) |
| CD 78 | + | + | + | + | PR11587 (251) |
| CD 79 | − | − | − | − | PR11528 (009) |
| CD 81 | − | − | − | − | PR1128 (009) |
| CD 82 | + | + | − | − | PR11664 (412) |
| CD 83 | + | + | − | − | PR11577 (449) |
| CD 85 | − | − | − | − | PR11693 (010) |
| CD 86 | − | − | − | − | PR11665 |
| CD 89 | + | + | + | + | PR11587 (251) |
| CD 90 | + | + | − | − | PR11666 (020) |
| CD 91 | − | − | − | − | PR11667 (593) |
| CD 92 (I) | + | + | − | − | PR11589 (106) |
| CD 92 (II) | − | − | − | − | PR11656 (039/2) |
| CD 92 (III) | + | + | − | − | PR11692 |
| CD 95 | + | + | − | − | PR11668 (020) |
| CD 96 | + | + | − | − | PR11577 (449) |
| CD 97 | − | − | − | − | PR 11528 (009) |
| CD 99 | − | − | − | − | PR11528 (009) |
| CD 100 | + | + | − | − | PR11666 (020) |
| CD 102 | − | − | − | − | PR11664 (412) |
| CD 104 | + | + | − | − | PR 11589 (106) |
| CD 112 | + | + | − | − | PR 13474 |
| CD 113 | + | + | − | − | PR11485 (001) |
| CD 115 (I) | − | + | − | − | PR11488 (017) |
| CD 115 (II) | − | − | − | − | PR11693 (010) |
| CD 116 | + | + | − | − | PR11685 (AI-82/1) |
| CD 117 (I) | + | + | − | − | PR11589 (106) |
| CD 117 (II) | + | + | − | − | PR11673 (AI-83) |
| CD 117 (III) | + | + | − | − | PR11674 (235) |
| CD 119 | + | + | − | − | PR11675 (AI-8/0) |
ID, identification.
Ribotypes that start with PR were the IDs given by Webribo database. The number in the parentheses is the accepted best match given by the same database. When the band pattern was the same and the differences of fragment sizes were ±4 bp, the most likely ribotype was accepted. Otherwise, it was considered a new ribotype.
ND, presence of tcdA or deleted tcdA was not observed.
Antimicrobial resistance.
All C. difficile isolates (n = 58) isolates were susceptible to vancomycin (MIC90, 1 μg/ml), metronidazole (MIC90, 2 μg/ml), linezolid (MIC90, 2 μg/ml), tigecycline (MIC90, 4 μg/ml), and moxifloxacin (MIC90, 4 μg/ml). However, a reduced susceptibility was observed for clindamycin in 53.45% of the C. difficile isolates (Table 4). When all the clindamycin-resistant isolates were tested for the ermB gene, only 4 isolates had positive bands, and all of those isolates had a breakpoint of 32 μg/ml for clindamycin, as determined by the agar dilution method. Furthermore, those four isolates were nontoxigenic and isolated from biosolid-based (n = 2), dairy manure-based (n = 1), and poultry litter-based (n = 1) composts.
TABLE 4.
Analysis of antimicrobial susceptibilities of C. difficile strains isolated from manure/compost samples by agar dilution method
| Antimicrobial | Breakpoint or MIC data (μg/ml)a |
Resistance (%)b | |||
|---|---|---|---|---|---|
| Breakpoint | MIC range | MIC50 | MIC90 | ||
| Metronidazole | ≥32c | ≤0.125–64 | <0.125 | 2.00 | ND |
| Moxifloxacin | ≥8c | ≤0.125–32 | 2.00 | 4.00 | ND |
| Clindamycin | ≥8c | ≤0.125–64 | 8.00 | 16.00 | 53.45 |
| Vancomycin | ≥16 | ≤0.125–32 | 0.25 | 1.00 | ND |
| Tigecycline | ≥8d | ≤0.125–16 | 1.00 | 4.00 | ND |
| Linezolid | ≥8d | ≤0.125–16 | 2.00 | 2.00 | ND |
MIC range, tested concentration range of antibiotics. MIC50 and MIC90 are the concentrations of antibiotics at which 50 and 90% of the isolates were inhibited, respectively.
ND, not detected.
Antimicrobial resistance breakpoints according to the CLSI (32).
Antimicrobial resistance breakpoints according to Rodriguez-Palacios et al. (9).
DISCUSSION
The study goal was to assess manure/compost as a potential carrier of the endospore-forming Gram-positive bacterium C. difficile. The compost samples collected for this study represented a wide variety of feedstocks currently being used for composting in the United States. Having positive results for all categories of compost samples reveals the environmental sources in which C. difficile could persist. The contamination could be from the contaminated feedstock, soil, workers, wildlife, or postprocessing of the compost.
The prevalence of C. difficile in fecal matter, such as anaerobically digested sludge from wastewater treatment plants, dewatered biosolids (13), and farm animal fecal matter (6), has been reported. The composition of feedstock, moisture content, and storage temperature affect the survival of many foodborne pathogens in animal manure and compost (14). In this study, significant correlations (P < 0.05) were detected between the presence of C. difficile endospores in manure/compost and the physical parameter, MC, and microbiological parameters, such as thermophile and mesoactinomycete counts. However, there are differences in the whole model and stepwise analysis, which could be due to either the natural trend or the small sample size (<30 samples in unprocessed manure) for a fair statistical analysis.
According to the information acquired from the compost makers, most of the composting processes were conducted with a regulated thermophilic phase, where the temperatures were >55°C (>131°F) for >3 days in order to comply with U.S. Environmental Protection Agency regulations (15) or USDA organic composting guidelines (16) to reduce the levels of most pathogenic bacteria in feedstocks. The lower level of fecal coliforms in most finished compost samples of the current study implies that the treatments used in composting eliminated most Gram-negative pathogens of Enterobacteriaceae family. Fecal coliforms are commonly used as the indicators of fecal contamination in environmental samples. However, in this study, the presence of fecal coliforms does not correlate with the presence of C. difficile, suggesting the difference between fecal coliforms and the resistant C. difficile endospores when exposed to composting temperatures. In agreement with that, Fujioka and Shizumura (17) reported that fecal coliforms are not reliable as an indicator in testing the fecal contaminations due to the inconsistent levels of contamination and confirmed that endospore-forming C. perfringens is a better indicator due to consistency of endospore concentration in drinking water.
Clostridia form heat-resistant endospores may survive the thermophilic phase of composting and certain cooking temperatures (18). Windrow composting was reported to reduce the majority of C. difficile endospores (>99%) effectively compared to the land application of dewatered biosolids (4). The prevalence of C. difficile in swine manure compost was assessed previously to be 36%, where piglet manure was a fraction of feedstock in all the samples (11). Previous studies reported that young animals, especially piglets, carry C. difficile more than adult pigs (19). At mesophilic temperatures, C. difficile endospores in swine manure were reported to germinate and proliferate in the population by 100-fold (11). Upon land application of contaminated compost, anaerobic environments in subsurface soil create suitable conditions for the proliferation of vegetative cells of clostridia (20). Therefore, these studies suggest that the compost could be a potential environment for the survival and proliferation of anaerobic and endospore-forming C. difficile strains as well.
In our study, 40% of the plant and food waste compost samples were positive for C. difficile. The isolation of C. difficile from raw meat samples (21) suggested that raw or undercooked meat was a potential source of C. difficile in food waste. Moreover, C. difficile was isolated from compost prepared solely of plant materials, too. One of the C. difficile-positive plant-based composts was made from plant leaves collected from curbs in a municipal area, which could possibly be contaminated with the infected pet animal feces.
The major typing method for C. difficile used in this study was capillary gel electrophoresis-based PCR-ribotyping (22). For some isolates, there were slight differences between the experimental and reference band patterns in the Webribo database. Indra et al. (22) explained that an error margin of ±4 bp existed in creating the database. Hence, the “most likely” match was accepted when the deviations were within ±4 bp of fragments. Even the two reference strains used in the validation were also not identified as ribotypes 060 and 251 directly, but the most likely matches were accurate for those two strains. When the database selection was most likely, the band pattern and DNA fragment sizes of the isolate were compared with the database match. For example, the isolate with A?B+CDT+ was identified as ribotype 078, and every fragment had a deviation of <0.5 bp compared to the ribotype 078 in the database. The differences in DNA fragment sizes observed in this study could be due to the use of different DNA extraction kits, PCR master mixes, genetic analyzer, column length, and polymer packed in the column from the ribotypes reported by other researchers in the database.
Toxigenic C. difficile strains harbor genes for the main virulence factors toxin A (tcdA), an enterotoxin, and toxin B (tcdB), a cytotoxin in the pathogenicity locus (PaLoc), which is a highly stable and conserved region in the chromosome (23). The presence of toxins in C. difficile can be used to classify toxigenic strains (A+B+), strains producing only toxin B (A−B+), and nontoxigenic strains (A−B−). Previous studies have demonstrated the increased diversity of C. difficile strains due to horizontal gene transfer and recombination in the PaLoc. These variations are commonly found among A+B+ strains from the reference strain, VPI 10463 (23). In this study, 25 A+B+ C. difficile ribotypes were observed among 38 isolates from manure/compost samples. Based on the literature, four clinically important A−B+ ribotypes of C. difficile, 036, 017, 047, and 110, have been detected (23) compared to only ribotype 017 (A−B+) present among our isolates, which has the deleted fragment of tcdA. The one isolate with undecided results for tcdA (A?B+CDT+) should have an altered sequence for tcdA or the primer binding site. The absence of deleted tcdA fragment is evidence for the existence of alterations in tcdA, and so its toxigenic profile cannot be determined as A−B+. The third toxin type in C. difficile, binary toxins, cause some cytotoxic effect in cell cultures (24) and increase the adherence of C. difficile to intestinal cells by destroying the actin cytoskeleton (25). Binary toxins are more prevalent among the variant A+B+ strains (24). In agreement with this, 2 out of 3 isolates with binary toxin genes detected in this study were positive for A+B+, and the third one was the isolate positive for tcdB but negative for both tcdA and deleted tcdA. The presence of binary toxins in nontoxigenic isolates had also been reported in the United States (24). However, no binary toxin-only strains were identified in the present study.
Toxigenic strains of C. difficile have previously been isolated from fresh retail vegetables (both root and aboveground vegetables) and ready-to-eat vegetable salads (7–10). A recent study by Rodriguez-Palacios et al. (9) isolated toxigenic C. difficile strains (3/125) from aboveground-grown vegetables sold in retail. Although the prevalence is low, some studies confirmed that vegetables also carry both toxigenic and antimicrobial-resistant C. difficile isolates (8–10). Some ribotypes (001, 014, 017, and 078) isolated from vegetables (7–10) were the same as the ribotypes determined in the present study from animal manure/compost samples; however, a definite conclusion on how vegetables were contaminated with C. difficile could not be drawn due to lack of information from pre- and postharvesting statuses. Therefore, future studies using more sensitive tracking tools will be able to identify the original sources of contamination of vegetables. As required by the Food Safety Modernization Act (FSMA), the FDA established standards to improve the safety of produce for human consumption under pre- and postharvest conditions by emphasizing the microbial safety of biological soil amendments, such as stabilized compost used in growing produce (26). Accordingly, stabilized compost should be used, and the potential for contact with produce should be minimized after application. However, Xu et al. (4) revealed that endospores of pathogenic C. difficile survived well in windrow composting of biosolid waste after an extended 47-day thermophilic phase of composting. Therefore, if the length of thermophilic phase of composting is shorter, those compost products could be a source of C. difficile in vegetables.
Dissemination of antimicrobial resistance (AMR) in the land application of compost contaminated with AMR strains has been suggested previously (27). Some phylogenetic groups of E. coli isolated from animal manure-based compost were identified by their carriage of resistance genes to multiple antibiotics (27). Although there is a little information on the antimicrobial resistance of C. difficile in compost (11), strains resistant to multiple antibiotics have been isolated from livestock fecal samples (28). AMR is detected in laboratories using breakpoints to explain the MIC of an antimicrobial agent that separates wild-type bacteria from the resistant bacteria (29, 30). Usually, the term “breakpoint” is used only if the data represent a therapeutic application, and the European Committee on Antimicrobial Susceptibility Testing (EUCAST) recommends the use of the term “epidemiological cutoff value” for nonclinical applications (29, 30). However, regarding all antimicrobials tested against C. difficile, there are no epidemiological cutoff values according to Clinical and Laboratory Standards Institute (CLSI) guidelines. The majority of our isolates (31/58) had reduced susceptibility to clindamycin, as observed using the agar dilution method.
Previous literature also reported the reduced susceptibilities of environmental and community-associated C. difficile isolates to clindamycin (28). Clindamycin resistance in C. difficile, a risk factor for CDI, is due to carriage of the ermB gene (31). In this study, only 4 nontoxigenic strains out of the 31 clindamycin-resistant isolates from the agar dilution method were positive for the ermB gene. The MIC of those 4 strains was recorded as 32 μg/ml (clindamycin breakpoint, 8 μg/ml) by the agar dilution method (32). The same discrepancy for clindamycin resistance was reported by Dong et al. (33) after analyzing clinical C. difficile isolates for antimicrobial susceptibilities by the agar dilution method and PCR detection of ermB. Low-level clindamycin resistance (8 to 24 μg/ml) in ermB-negative isolates has been observed due to a point mutation in this gene (34). Although the previous studies reported the clindamycin resistance of C. difficile strains isolated from different environment samples, it is difficult to compare our results directly to those of these studies, as the methods used in assessing the antimicrobial resistance were different from each other (9, 28). Due to the presence of many mobile genetic elements in the genome of C. difficile, nontoxigenic strains may become toxigenic, and antimicrobial-susceptible strains may become antimicrobial resistant by horizontal gene transfer (35). Therefore, clindamycin resistance in nontoxigenic strains of C. difficile should not be overlooked.
Limitations.
In this study, the samples were collected according to the feasibility of collection and positive responses from the animal waste processers to our requests for samples. Therefore, the results reported here may not completely represent the prevalence of C. difficile in compost produced in the United States due to the limited number of samples examined. Moreover, the age of the animal is important in considering the contaminations of animal manure with C. difficile, as the pathogen has been isolated more frequently in the fecal matter of young animals. In this study, the data on the age of the animals that produced manure were not available. A larger survey of compost products is needed to conclude the prevalence and the factors that influence the persistence of C. difficile in finished compost in the United States. Analysis of samples from composting from different feedstocks and at selected time intervals of the composting process will provide scientific evidence on the sources of contamination and the fate of C. difficile endospores at each phase of composting. Since a positive sample obtained by an enrichment method may be due to a single viable spore, further studies that enumerate the population of C. difficile endospores in manure/compost are recommended to accurately reflect the potential of those environmental samples as sources of C. difficile.
Conclusion.
This study demonstrated that 36% of the composts used in agroproduction were contaminated with C. difficile. Even though the foodborne transmission of C. difficile has not been clearly established, compost contaminated with C. difficile could likely transmit the pathogen to the edible parts of the fresh produce, when it is used as a biological soil amendment. The results of this study indicate that not only unprocessed animal manure, but also properly processed finished composts, are a reservoir of toxigenic C. difficile ribotypes, such as 106, 020, 412, and 251. Our study revealed that antimicrobial resistance is rare among those C. difficile isolates obtained from compost. However, clindamycin resistance may be disseminated via compost. Further studies on potential contamination of fresh produce by pathogenic C. difficile via compost need to be conducted. Additionally, current regulations on composting might need to be reevaluated regarding the survival of endospore-forming pathogens in manure/composts.
MATERIALS AND METHODS
Sample collection.
A total of 142 manure and compost samples of different feedstocks were collected from multiple states in the United States, including California, Colorado, Delaware, Florida, Georgia, Illinois, Iowa, Kentucky, Massachusetts, Michigan, Nevada, New Mexico, New York, Pennsylvania, South Carolina, South Dakota, Tennessee, Texas, Vermont, Virginia, Washington, and Wisconsin. Out of those samples, 113 samples were collected between 2014 and 2015. Twenty-one unprocessed manure samples were collected, and most of them were from the farms in the vicinity of Clemson, SC, from sources including chicken/turkey litter (n = 12), cow manure (n = 2), and mixed animal manure (n = 7). Finished commercial compost samples (n = 91) were requested from the compost makers, and an additional sample was purchased from a local supplier. Samples were collected in Ziploc bags (S. C. Johnson & Son, Racine, WI), shipped under ambient conditions, and stored under refrigeration conditions (4°C) upon arrival until analysis. Additionally, 29 finished compost samples collected in 2011 and 2012 and stored at −10°C from our previous study (36) were analyzed.
Physical and microbiological analysis.
The water activity of each sample was measured using a dewpoint water activity meter (AquaLab series 3TE; Decagon Devices, Pullman, WA), and the moisture content (MC) was determined using a moisture analyzer (model IR-35; Denver Instrument, Denver, CO). The pH values manure/compost were measured by a multiparameter benchtop meter (Orion Versa Star meter; Thermo Fisher Scientific, Inc., Fort Collins, CO) according to the test methods described by the U.S. Composting Council (37).
Selected dilutions from the prepared 10-fold serial dilutions were plated on tryptic soy agar (TSA; Becton & Dickinson, Sparks, MD), actinomycetes agar (Becton & Dickinson), and Rose-Bengal agar (CM 0549; Oxoid, Lenexa, KS) supplemented with chloramphenicol (100 mg/liter; SR 0078; Oxoid) to enumerate bacteria, actinomycetes, and yeast/molds, respectively. Incubation temperatures of 37, 55°C, and room temperature (∼22°C) were used for mesophiles and mesoactinomycetes, thermophiles and thermoactinomycetes, and yeast/molds, respectively. Coliforms and E. coli were enumerated by plating 1 ml of the selected dilutions on E. coli/coliform Petrifilm (3M, St. Paul, MN) and incubating overnight at 37°C. All samples were tested in duplicate.
Isolation of C. difficile from manure and compost samples.
Qualitative detection of C. difficile from manure/compost samples was performed using brain heart infusion enrichment broth (Becton & Dickinson) supplemented with moxalactam (32 mg/liter; Alfa Aesar, Haverhill, MA), norfloxacin (12 mg/liter; Alfa Aesar), l-cysteine (1 g/liter; Alfa Aesar), and sodium taurocholate (1 g/liter; Alfa Aesar), as previously described (36). Briefly, 5 g of each sample was added to 20 ml of enrichment broth in a sterile 50-ml conical tube (Corning, Tewksbury, MA), mixed by vortexing, and then incubated anaerobically at 37°C for 7 days with loose lids inside an anaerobic jar with anaerobic gas packs (Becton & Dickinson). Following the incubation, 1 ml of each sample was heat-shocked at 60°C for 25 min in a water bath (model no. DL 30; Haake, Paramus, NJ) and centrifuged at 4,000 × g for 10 min. The supernatant was discarded, and the pellet was resuspended in 50 μl of sterile distilled water and streaked onto C. difficile agar supplemented with 7% horse blood (Remel, Lenexa, KS), sodium taurocholate (1 g/liter), l-cysteine (1 g/liter), moxalactam (32 mg/liter), norfloxacin (12 mg/liter), and cycloheximide (50 mg/liter). After the anaerobic incubation of plates at 37°C for 24 h, presumptive positive colonies were identified by nonhemolytic, swarming, flat-colony morphology, Gram staining, and l-proline aminopeptidase activity using PRO discs (Remel).
PCR analysis for the identification of C. difficile.
Bacterial genomic DNA was extracted using a microbial DNA extraction kit (Mo Bio, Carlsbad, CA), and the extracted DNA was stored at −20°C until use. Presumptive C. difficile isolates were confirmed by touchdown PCR (38) for detection of the species-specific housekeeping gene tpi (39), using previously published primers (Table 5), and C. difficile ATCC 43593 served as the positive control. All PCRs were performed in an Eppendorf thermal cycler (Mastercycler Realplex2; Eppendorf, Westbury, NY) in a final volume of 25 μl/reaction mixture, which consisted of 10 mM amplification buffer, 2 mM MgCl2, 0.2 mM each deoxynucleoside triphosphate, 0.4 μM each primer, 1 unit of Taq polymerase (TaKaRa Bio, Inc., Japan), and 2 μl of template DNA. PCR-positive isolates were further confirmed by a latex agglutination test for C. difficile (DR 1107; Oxoid).
TABLE 5.
PCR primers used in this studya
| Gene or ribotyping | Primer sequence (5′ to 3′) (primer name) | Product size (bp) | Reference |
|---|---|---|---|
| tpi | AAAGAAGCTACTAAGGGTACAAA | 230 | 39 |
| CATAATATTGGGTCTATTCCTAC | |||
| tcdA | GATTCCTATATTTACATGACAATAT | 369 (undeleted) | 39 |
| GTATCAGGCATAAAGTAATATACTTT | 110 (deleted) | ||
| tcdB | GGAAAAGAGAATGGTTTTATTAA | 160 | 39 |
| ATCTTTAGTTATAACTTTGACATCTTT | |||
| cdtA | TGAACCTGGAAAAGGTGATG | 376 | 40 |
| AGGATTATTTACTGGACCATTTG | |||
| cdtB | CTTAATGCAAGTAAATACTGAG | 510 | 40 |
| AACGGATCTCTTGCTTCAGTC | |||
| ermB | AATAAGTAAACAGGTAACGTT (primer 2980) | 688 | 31 |
| GCTCCTTGGAAGCTGTCAGTAG (primer 2981) | |||
| Ribotyping | |||
| 16S | FAM-GTGCGGCTGGATCACCTCCT | Variable | 41 |
| 23S | CCCTGCACCCTTAATAACTTGACC |
Primers for the 16S and 23S rRNA regions are shown for ribotyping. The 16S primer is capped with 6-carboxyfluorescein (FAM) and corresponds to bases 1482 to 1501 of the 16S ribosomal gene, and the 23S primer corresponds to bases 1 to 24 of the 23S ribosomal gene.
Detection of toxin genes.
Genes encoding toxins A (tcdA) and B (tcdB) of C. difficile were detected using the primers described previously (39) (Table 5). Genes for binary toxins were detected by screening the genomic DNA for cdtA and cdtB genes, as previously described (40) using C. difficile ATCC BAA 2155 as the positive control. PCR products (5 μl each) were resolved on a 1.5% agarose gel for 60 min at 75 V and visualized after staining with ethidium bromide (0.5 μg/ml).
Capillary gel electrophoresis-based PCR-ribotyping.
PCR-ribotyping was performed using primers (41) (Table 5) and methods (22) described previously. PCR amplicons were purified using the UltraClean GelSpin PCR clean-up kit (Mo Bio) and analyzed using a genetic analyzer (model no. 3730xl; Applied Biosystems, Foster City, CA) with a t50-cm capillary loaded with a POP4 gel (Applied Biosystems). Samples were injected at 5 kV over 5 s with a total running time of 30 min at 15 kV run voltage. GeneScan 1200 LIZ standard (Applied Biosystems) was used as an internal marker. Fragment analysis results were obtained as .fsa files and uploaded to the Web-based database Webribo (http://webribo.ages.at) to identify if C. difficile ribotypes distributed in manure/composts were outbreak strains. C. difficile ATCC strains 43593 (ribotype 060) and BAA 2155 (ribotype 251) were used to validate ribotyping analysis.
Antimicrobial resistance.
All confirmed C. difficile isolates were tested for susceptibility to the selected antimicrobials: metronidazole (MT), vancomycin (VN), moxifloxacin (MX), clindamycin (CL), linezolid (LN), and tigecycline (TG) (Sigma-Aldrich, St. Louis, MO) using the agar dilution method (32). Brucella blood agar (Himedia, Mumbai, India) supplemented with 5 μg/ml vitamin K (Sigma-Aldrich), 1 μg/ml hemin (Sigma-Aldrich) (32), and l-cysteine (1 mg/ml) was used as the culture medium. After C. difficile isolates were grown anaerobically on brucella blood agar for ∼24 h, colonies were suspended in 0.85% saline, and the turbidity of each suspension was adjusted to McFarland 0.5 (42). The suspensions were delivered to the supplemented brucella blood agar plates using a sterile 48-pin replicator (3 mm in diameter) (Sigma-Aldrich) aseptically. C. difficile ATCC 700057 and Bacteroides fragilis ATCC 25285 were used as quality control microorganisms. The inoculated plates, including the control plates, were incubated at 37°C for 48 h anaerobically, and 2 additional control plates were incubated at the same temperature aerobically to assess any aerobic contaminations in the suspensions prepared. Brucella blood agar plates for TG and LN concentrations were prepared on the same day of inoculation, whereas the plates for other antibiotic concentrations were prepared 1 to 2 days in advance. The epidemiological resistance (R) cutoff values for antimicrobials (metronidazole, ≥32 μg/ml; vancomycin, ≥4 μg/ml; tigecycline, ≥8 μg/ml; linezolid, ≥8 μg/ml; clindamycin, ≥8 μg/ml; moxifloxacin, ≥8 μg/ml; and chloramphenicol, ≥32 μg/ml) were used as described previously (9, 32) to determine the antimicrobial resistance. Additionally, C. difficile isolates with clindamycin resistance were further screened for the presence of the corresponding gene, ermB, by PCR as described previously (31) (Table 5).
Statistical analysis.
Bacterial counts on different plating media were compared by converting plate counts into log values as log CFU per gram of compost on the dry weight basis. Logistic regression of the Statistical Analysis System (JMP Pro 12; SAS Institute, Inc., Cary, NC) was used to reveal any correlations between the presence of C. difficile with the physical and microbiological factors tested. A complete model was used to determine the effect of any factor on the presence of C. difficile in compost/manure, and stepwise approaches were used to confirm the impact of each selected factor on the whole model in finished composts and unprocessed manure (P = 0.05).
ACKNOWLEDGMENTS
We thank William C. Bridges, Jr. (Clemson University), for assisting in the statistical analysis of data and Annel Greene (Clemson University) for critically reviewing the manuscript.
This research was partially supported financially by the Center for Produce Safety, University of California, Davis, and the U.S. Department of Agriculture (USDA) Agricultural Marketing Service through grant 15-SCBGP-CA-0046.
The contents of this article are solely the responsibility of the authors and do not necessarily represent the official views of the USDA.
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