Crocodiles are 1 of the few species that are long-lived and can endure environments that would otherwise be detrimental to human health and well-being.1 For example, crocodiles have evolved in harsh conditions, including unsanitary environments, consume rotten meat, are typically exposed to heavy metals such as arsenic, cadmium, chromium, cobalt, nickel, etc., and are also exposed to high quantities of solar radiation. Recently, we have speculated that species such as crocodiles can prosper in these environments, possibly due to their gut microbiome.2,3 This is not a novel concept as gastrointestinal tracts are unique environments inhabited by microbial flora over ancient evolutionary processes and have an important role in the health and disease of its host.4–14 Despite advances in analytical tools, a complete understanding of host-microbial interactions especially in reptiles such as crocodiles has remained incompletely understood. As bacterial colonization of the gut starts with the oral cavity, it is puzzling that very few studies have been conducted on the oral microbiome, especially in Crocodylus porosus. Among the few investigations of the oral cavity and the colon of the Alligator mississippiensis, the results revealed that the oral microbiome of the alligator was dominated by Proteobacteria, Bacteroides, and Firmicutes, while the colon was dominated by Bacteroides, Firmicutes, and Proteobacteria.15 This is in contrast to Kieran et al.16 which observed predominance of Firmicutes and Bacteroidota across all tissue types. It is important to compare the oral vs the intestinal tract to have a complete understanding of the C. porosus microbiome. Of note, in alligators, Keenan et al.5 showed that their gut microbiome comprised of Fusobacteria. More recently, Murphy et al.17 showed an abundance of Proteobacteria followed by Firmicutes in the gut of the American alligator. In the Australian saltwater crocodile (C. porosus), Willson et al.18 showed that the dominant phyla are Firmicutes. In this study for the first time in C. porosus, using 16S rRNA sequencing, bacterial diversity was compared in the oral cavity, the small intestine, and the large intestine.
Materials and Methods
C. porosus bacteria
Isolation and identification of crocodile bacteria from the gastrointestinal tract were accomplished as described previously.19 In brief, approval for the study was obtained from the Malaysian Department of Wildlife and National Parks (PERHILITAN), as well as Sunway University, Malaysia (SUNREC 2019/023). A convention on international trade in endangered species (CITES) of wild fauna and flora registered crocodile farm, provided the saltwater crocodile, C. porosus. Management of crocodile including anesthesia and dissection of the internal organs were carried out by qualified personnel at the farm who routinely perform these procedures. In brief, the crocodile was sedated using diazepam (0.5 mg/kg), administered intramuscularly (IM) in the hind limb using a pole syringe, and then ketamine at 40 mg/kg IM was used for about 3 hours duration, and euthanasia was performed via destruction of the brain function by instantaneously severing the spine behind the head and pithing. Next, the ventrum was cleaned with surgical antiseptics, and 1 long ventral incision was performed, the esophagus was cut above the stomach, and the large intestine and the entire gastrointestinal tract were removed onto an adjacent clean table. Then each incision was done with a new scalpel and the swabs were taken. Additionally, we also confirmed that all the experiments were carried out in agreement with appropriate protocols and guidelines as formerly defined.19 The whole gut was removed aseptically and bacteria from the gastrointestinal tract (the oral cavity, proximal region of the small intestine [jejunum], and distal part of the large intestine) were isolated using sterile cotton swabs, and 3 samples were taken from each segment of the gastrointestinal tract of a single animal (male).19 Crocodiles were held in captivity and fed on chicken. Crocodile was not fed any meal for 72 hours before experimentation. Three samples were taken from the oral cavity (directly from the gingiva), 3 samples from a similar region of the small intestine, and 3 samples from a similar region of the large intestine were obtained using sterile cotton swabs (tissues were of normal histology). Next, swabs were placed in 15 mL sterile centrifuge tubes, individually and kept on ice for immediate transportation to the laboratory. Bacterial identification was accomplished using 16S rRNA gene amplification and sequencing as described below.
Extraction of DNA
Amplicon sequencing investigations were carried out using DNA extracted from the gastrointestinal tract (the oral cavity, the small intestine, and the large intestine).20 The sample preparation and DNA extraction from the animal tissues have been described in our earlier study.21 Briefly, the samples were treated with 1 mL of heated lysis solution (comprising 20 g of SDS per liter, 0.1 M Tris-HCl, 0.15 M sodium chloride, and 25 mM EDTA). After the addition of 10 µM proteinase K (10 mg/mL), samples were incubated for 60 min at 65°C, and then centrifuged for 10 minutes at 12,000 X g, the residue was extracted using phenol, chloroform, and isoamyl alcohol in the following ratios: 25:24:1, 24:1, and 25:24:1 respectively. Utilizing potassium acetate (3 M, pH 5.5) and 95% ethanol, the DNA was deposited, and centrifuged for 10 minutes at 15,000 X g. This was followed by washing twice with 70% ethanol and mixed in 0.4 mL of buffer (10 mM Tris, 1 mM EDTA). To eliminate any remaining RNA, samples were treated for 30 minutes with 10 mg of RNAse enzyme at 37°C. Next, the protein was extracted using isoamyl alcohol and chloroform. The upper layer was removed from the DNA and placed in an Eppendorf tube comprising 2.5 vol of ethanol. The DNA was purified and spun at 15,000 X g for 10 minutes (second purification). The DNA was then resuspended in 0.2 mL of dH2O, and washed twice. The DNA content and purity were assessed with a 1% agarose gel.
16S rRNA sequencing
The 16S rRNA/ITS gene in different locations (V3-V4/16S) was amplified using specific primers (341F-CCTAYGGGRBGCASCAG, and 806R-GGACTACNNGGGTATCTAAT), as described elsewhere.20 Cycling conditions comprised a primary denaturation step at 98°C for 1 minute, ensued by 30 cycles at 98°C (10 seconds), 50°C (30 seconds), and 72°C (30 seconds), and a final 5 minutes extension at 72°C as described in detail previously.20 PCR products were combined with 1X loading buffer, containing SYB green, and electrophoresed on a 2% agarose gel.19 PCR products (470 bp) were purified using the Qiagen Gel Extraction Kit (Qiagen). NEBNext® UltraTM IIDNA Library Kit was utilized to generate the libraries (Cat No. E7645). Finally, 250 bp paired-end reads from the library’s sequencing on the Illumina NovaSeq platform were produced.
Data processing
The data were processed and effective tags (shown in Table 1) for subsequent analysis were obtained, as previously described.20 In summary, reads were truncated by trimming out primer sequences and barcodes and merged using FLASH (version 1.2.11, http://ccb.jhu.edu/software/FLASH/, accessed on January 15, 2023) to merge paired-end reads. The tags were compared with the reference database (Silva database https://www.arbsilva.de/ for 16S/18S, accessed on March 20, 2023, and Unite database https://unite.ut.ee/ for ITS, accessed on March 20, 2023) using Vsearch (version 2.15.0) and denoise was performed with DADA2 in the QIIME2 software (version QIIME2-202006) and multiple sequence alignment was performed using QIIME2. The OTUs produced with 97% sequence identity were regarded as homologous. When the clustering threshold was 97%, the following statistical indices of alpha diversity were calculated: the number of reads selected for normalization: cutoff = 12,466. Alpha diversity was calculated from indices in QIIME2 including Chao Index, Simpson Index, Shannon Index, ACE Index, and Good’s Coverage Index, and beta diversity was calculated in QIIME2 (such as unweighted unifrac and weighted unifrac distance). For differences in community structure between groups, adonis and anosim in QIIME2 were used. For different species at each taxonomic level, R software (version 3.5.3) was used to do MetaStat and t test analysis.
Summary of amplicon sequencing to generate 250 bp paired-end raw reads, and then merged, treated to obtain effective tags for subsequent analysis.
| Sample name | Raw PE reads (original reads) | Raw tags (merged Tags) | Clean tags (post-filtering) | Effective tags (post-filtering chimera) | No. of bases of effective tags | Average length of effective tags |
|---|---|---|---|---|---|---|
| A | 188,056 | 176,455 | 171,585 | 126,613 | 54,105,181 | 427 |
| B | 176,323 | 147,659 | 139,458 | 134,649 | 57,528,392 | 427 |
| C | 175,779 | 163,992 | 159,527 | 145,111 | 61,168,195 | 422 |
Results
Interspecific varieties in bacterial gastrointestinal communities
To determine bacterial flora variation in the gastrointestinal tract of C. porosus, 16S rRNA sequencing was performed as described in the Materials and Methods. Diversity was indicated by the rarefaction curve, and the OTUs in the sample were arranged according to their relative abundance using the rank abundance curve (or the number of sequences included). According to the findings, there were 153 species found in the oral cavity of C. porosus, 239 in the small intestine, and 119 in the large intestine (Figure 1). These findings suggest that the small intestine reflects the highest richness of bacterial communities compared with the oral cavity. In contrast, the large intestine exhibited the least richness of microbial communities in the gastrointestinal tract compared with either the oral cavity or the small intestine (Figure 1). Overall, the bacterial diversity was greater in the small intestine, while the large intestine exhibited relatively less microbial diversity.
(i) Rarefaction curve indicating the richness of microbial community in the gastrointestinal tract including, oral cavity (A), small intestine (B), and large intestine (C). (ii) Venn diagram showing the distribution of bacterial operational taxonomic units (OTUs) in the gastrointestinal tract including, oral cavity (A), small intestine (B), and large intestine (C), using sterile cotton swabs, according to the 16S rRNA gene sequence analysis as described in Materials and Methods section.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.03.0061
As shown (Figure 1), various samples of the gastrointestinal tract of C. porosus included diverse bacterial OTUs, including the oral cavity (A), the small intestine (B), and the large intestine (C) indicating bacterial diversity. It is noteworthy that 156 OTU communities were specifically present in the small intestine (B), while 72 OTUs were specifically present in the oral cavity (A). Additionally, 57 OTUs were specific to the large intestine (C; Figure 1). We observed 9 OTUs shared between the small intestine (B), and the large intestine (C); 28 OTU communities shared between the oral cavity (A) and the small intestine (B); and 7 OTUs shared between the oral cavity (A) and the large intestine (C). On the other hand, all groups exhibited common bacterial OTUs in 46 intra-section regions. Overall, the results of the OTU analysis revealed both common and unique OTUs in the oral cavity, the small intestine, and the large intestine, with the small intestine showing the highest level of bacterial community richness.
Relative abundance of taxa on the level of phyla
Next, we determined the relative abundance of bacterial taxa. The relative abundance of taxa with comparative taxa with a high relative abundance is depicted (Figure 2), as is the proportion of each sample at different taxonomic levels. Annotation data from the taxa depicts the top 10 taxa at the various taxa levels (phylum, class, order, family, and genus). The relative abundances of taxa within phyla are displayed (Figure 2). The term “Other” denotes all phyla save the top 10 phyla collectively. Interestingly, the most common ph“lum w”s Proteobacteria, which was identified throughout the oral cavity (A) and the gastrointestinal tract (the small intestine (B) and the large intestine (C).
Distribution of dominant phyla (i) and genus (ii) in the gastrointestinal tract including, oral cavity (A), small intestine (B), and large intestine (C).
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.03.0061
Overall, the relative abundance in the oral cavity of C. porosus was in the order of Proteobacteria, Bacteroidota, Firmicutes, Actinobacteriota, Synergistota, Chloroflexi, Desulfobacterota, and others. Meanwhile, the relative abundance in the small intestine of C. porosus was in the order of Proteobacteria, Bacteroidota, Firmicutes, Actinobacteriota, Synergistota, Desulfobacterota, Chloroflexi, and others. However, the relative abundance in the small intestine of C. porosus was in the order of Proteobacteria, Bacteroidota, Firmicutes, Actinobacteriota, Desulfobacterota, Synergistota, Chloroflexi, and others.
Taxa relative abundances at genus level
The abundance of bacteria at the genus level in the gastrointestinal tract of C. porosus was determined. At the genus level, the relative abundance of bacteria in the oral cavity of C. porosus was in the order of Pseudomonas, Stenotrophomonas, Castellaniella, Comamonas, Alishewanella, Achromobacter, Fluviicola, Sphingobacterium, Pannonibacter, Salmonella, and others (Figure 2). While in the small intestine of C. porosus the relative abundance was in the order of Pseudomonas, Comamonas, Salmonella, Stenotrophomonas, Pannonibacter, Castellaniella, Achromobacter, Alishewanella, Sphingobacterium, Fluviicola, and others (Figure 2). In contrast, the relative abundance of bacteria in the large intestine of C. porosus was in the order of Salmonella, Pannonibacter, Comamonas, Pseudomonas, Stenotrophomonas, Castellaniella, Alishewanella, Achromobacter, Fluviicola, Sphingobacterium, and others (Figure 2). In short, these results showed that the distribution of the genus Pseudomonas between the oral cavity and the large intestine was different. Both the oral cavity and the small intestine had the same ratio of the genus Pseudomonas in contrast to the large intestine, which had less distribution of the genus Pseudomonas. The relative abundance of Stenotrophomonas, Castellaniella was increased in the oral cavity compared with the small intestine, while the relative abundance of Comamonas, Salmonella was increased in the small intestine compared with the oral cavity. In contrast, the relative abundance of Salmonella, and Pannonibacter was greater in the large intestine.
Analysis of the Principal Component (PCA) and Principal Coordinate (PcoA)
Bacterial communities in different samples were compared using Beta Diversity analysis, and the dissimilarity between samples was estimated as a square matrix of “distance” or “dissimilarity.” Principal Component Analysis (PCA) and Principal Coordinate Analysis (PCoA) were used to visualize the data in this distance matrix. How the principal component analysis (PCA) revealed differences in the bacterial diversity between the oral cavity, the small intestine, and the large intestine is illustrated (Figure 3). When main factors PC1 and PC2 were evaluated, the findings showed that PC1 contributed 61.18% while PC2 contributed 38.82%. Overall, these findings show that the groups’ bacterial communities exhibited differences. Clustering analysis was used to create a cluster tree to examine the similarity between various data. A phylogenetic tree was developed from a distance matrix utilizing the Unweighted Pair-group Method with Arithmetic Mean (UPGMA). To study the similarity among different samples, clustering analysis was used to construct a cluster tree. The results after integrating the clustering outcomes with the relative abundance of each sample per phylum are illustrated (Figure 4).
(i) Principal Component Analysis (PCA) and (ii) Principal Coordinate Analysis (PCoA) for the differences in microbial communities between the oral cavity (A), small intestine (B), and large intestine (C) of the C. porosus.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.03.0061
Unweighted Pair-group Method with Arithmetic Mean (UPGMA) cluster tree based on Weighted Unifrac distance between microbial communities of the oral cavity (A), small intestine (B), and large intestine (C) of the C. porosus.
Citation: American Journal of Veterinary Research 84, 8; 10.2460/ajvr.23.03.0061
Discussion
For the first time, in a single C. porosus, we compared bacterial diversity in the oral cavity, the small intestine, and the large intestine using 16S rRNA sequencing. In this study, we used a captive male C. porosus, at one time of its reproductive activity, on a singular diet, hence this cannot be taken as generalized for all C. porosus. With this limitation in mind, our study revealed that Proteobacteria, Bacteroidetes, and Firmicutes were dominant in the oral cavity, the small intestine, and the large intestine; however, differences were observed at the genus level, especially related to the oral cavity and the large intestine, albeit these are qualitative findings and further research is needed to substantiate these conclusions. These findings clearly demonstrate that bacterial flora varies throughout the gastrointestinal tract of a healthy crocodile. Given that the microbiome composition plays an important role in the host physiology and well-being, these findings may form the basis of constituting microbiome as a potential biomarker in determining the overall health and well-being of C. porosus, as well as determining feed additives such as probiotics; however, intensive further research is needed to realize these expectations,
In our findings, the small intestine exhibited the highest bacterial diversity, which may potentially prevent intestinal dysbiosis and its systemic consequences. These are interesting and encouraging results. Although the overall trend of bacterial presence at the phyla level remains consistent with earlier studies carried out in other reptiles (Proteobacteria, Bacteroidetes, Firmicutes), however, some differences were observed especially at the species level. This could be due to differences in the reptilian species, their habitat, the environment, or diet (chicken in our study), or geographical location and this needs further investigation. Notably, this is the first such study to be carried out on C. porosus in Malaysia (Langkawi) which may also explain differences. Prospective studies should explore variations in the diet of the crocodiles in captivity, their age, sex, and time of the year may all play a role in their microbiome composition. Furthermore, examination of tissue samples, as well as their exposure to chemicals such as anti-parasitics, anti-coccidial feed, etc. should be considered. In the future, large-scale studies are needed to compare and validate these findings. Notably, our earlier studies have identified potent antimicrobial and anti-tumor effects of gut bacterial metabolites of C. porosus,1,22 hence a complete understanding of C. porosus microbiome has the potential of yielding novel bioactives of medicinal value. Prospective studies are necessitated to comprehend the precise role of the microbiome and the detailed mechanisms of the metabolites from the gut bacteria. In addition, comprehension of the microbiome to the ability of their host in enduring harsh conditions, as well as demonstrate increased longevity and little or reduced cellular senescence, and the overall health of C. porosus is required.
Acknowledgments
No external funding was used in this study. The authors declare that there were no conflicts of interest.
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