Intestinal Microbiome Confers Strong Colonization Resistance Against Necrotic Enteritis

Jing Liu 1

Jiaqing Guo 1

Isabel Tobin 1

Melanie A. Whitmore 1

Dohyung M. Kim 1

Prasiddha Paudel 1

Anisha Subedi 1

Michael G. Kaiser 2

Susan J. Lamont 2

Guolong Zhang 1,3,4,5✉ Emailzguolon@okstate.edu

Department of Animal and Food Sciences Oklahoma State University Stillwater Oklahoma USA

2 Department of Animal Science Iowa State University Ames Iowa USA

3 Department of Microbiology and Molecular Genetics Oklahoma State University Stillwater Oklahoma USA

4 Department of Biochemistry and Molecular Biology Oklahoma State University Stillwater Oklahoma USA

5 Department of Veterinary Pathobiology Oklahoma State University Stillwater Oklahoma USA

Jing Liu^1,†, Jiaqing Guo¹, Isabel Tobin¹, Melanie A. Whitmore^1,‡, Dohyung M. Kim¹, Prasiddha Paudel¹, Anisha Subedi¹, Michael G. Kaiser², Susan J. Lamont², and Guolong Zhang^1,3,4*

¹ Department of Animal and Food Sciences, Oklahoma State University, Stillwater, Oklahoma, USA

² Department of Animal Science, Iowa State University, Ames, Iowa, USA

³ Department of Microbiology and Molecular Genetics, Oklahoma State University, Stillwater, Oklahoma, USA

⁴ Department of Biochemistry and Molecular Biology, Oklahoma State University, Stillwater, Oklahoma, USA

Running Title: Colonization Resistance Against Necrotic Enteritis

^* Correspondence: Guolong Zhang, zguolon@okstate.edu

^† Current address: IEH Laboratories and Consulting Group, Lake Forest Park, Washington, USA

^‡ Current address: Department of Veterinary Pathobiology, Oklahoma State University, Stillwater, Oklahoma, USA

ABSTRACT

Necrotic enteritis (NE), caused by Clostridium perfringens, is a major enteric disease in poultry that leads to severe dysbiosis, morbidity, and mortality. Modulating the intestinal microbiota holds promise for enhancing animal health and disease resistance; however, specific commensal bacteria associated with NE protection remain elusive. Chicken breeds differ markedly in disease susceptibility, with Fayoumi chickens exhibiting greater resistance than Leghorn and Cobb chickens. We hypothesized that Fayoumi chickens harbor unique commensal bacteria that confer robust colonization resistance against NE. To test this, we challenged two inbred lines, Fayoumi M5.1 and Leghorn Ghs6, alongside commercial Cobb broilers with NE. Among these, M5.1 chickens demonstrated the highest resistance to NE. Cecal microbiota transplantation from the three breeds into newly hatched Cobb chicks revealed that M5.1-derived microbiota provided completion protection against NE. Comparative microbiome analysis demonstrated significant differences among breeds under both healthy and NE-challenged conditions. Notably, Bifidobacterium, largely absent in healthy chickens of all three breeds, was highly enriched in both the ileum and cecum of M5.1 chickens following NE challenge. Furthermore, oral administration of Bifidobacterium pseudolongum significantly reduced NE mortality in Cobb chickens. Collectively, these findings highlight the protective role of commensal bacteria from NE-resistant Fayoumi chickens and suggest their potential for microbiota-based strategies to mitigate NE in poultry.

KEYWORDS:

Fayoumi chickens

poultry

cecal microbiota transplantation

microbiota

colonization resistance

necrotic enteritis

Clostridium perfringens

Bifidobacterium

Lactobacillus

Ligilactobacillus salivarius

INTRODUCTION

Necrotic enteritis (NE) is an enteric disease caused by Clostridium perfringens, a Gram-positive, anaerobic, spore-forming, rod-shaped bacterium. NE remains one of the most economically devastating diseases in the global poultry industry, costing over $6 billion annually in production losses ¹. Various predisposing factors are known to promote NE development, such as exposure to apicomplexan protozoa, Eimeria, the etiologic agent of avian coccidiosis ². Traditionally, prophylactic in-feed antibiotics have proven effective for the control of clostridial infections in chickens ³. However, with recent withdrawal of in-feed antibiotics from livestock production in a growing number of countries and increasing consumer demand for antibiotic-free animal products, NE has become more prevalent, necessitating effective antibiotic alternatives ⁴. Probiotics such as Lactobacillus, Enterococcus, and Bacillus and prebiotics such as mannan-oligosaccharides have proved beneficial in NE alleviation ^3,4. However, no commensal bacteria have been directly linked to NE resistance, and NE-resistant commensals could potentially offer prophylactic and/or therapeutic utility against NE.

Two highly inbred chicken lines, Fayoumi M5.1 and Leghorn Ghs6, are drastically different in their ability to resist Newcastle disease, with M5.1 being highly resistant and Ghs6 highly susceptible ^5,6. Additionally, Fayoumi chickens are known to be more resistant to coccidiosis, avian influenza, and salmonellosis than Leghorns or commercial broiler chickens ^7–10. The intestinal microbiota plays a significant role in host homeostasis and health, and there is mounting evidence indicating that the microbiota is shaped by host genetics ^11–13. Thus, we hypothesize that the intestinal microbiota from disease-resistant chicken breeds harbors a unique microbiota that offers strong colonization resistance (CR) to diseases such as NE.

To test our hypothesis, we assessed the efficacy of the microbiota from M5.1, Ghs6, and Cobb chickens in protecting newly-hatched Cobb chickens from NE through cecal microbiota transplantation (CMT) and revealed that M5.1 microbiota is the most protective against NE. We further compared the intestinal microbiota profiles among the three chicken breeds under both healthy and NE conditions and identified a large number of differentially enriched bacteria. Additionally, we demonstrated that oral administration of Bifidobacterium pseudolongum, which was specifically enriched in NE-challenged M5.1 chickens, confers strong protection to Cobb chickens against NE challenge. These findings provide direct evidence that the intestinal microbiota, particularly from disease-resistant chickens, offers robust CR against NE and can be leveraged for NE mitigation.

RESULTS

Differential susceptibility of three chicken breeds to NE

To investigate the difference in NE susceptibility, we challenged M5.1, Ghs6, and Cobb chickens to induce NE (Fig. 1A). While approximately 80% of Cobb chickens and 91% Ghs6 chickens survived at 3 days post-infection (dpi), 100% M5.1 chickens exhibited no mortality (Fig. 1B). Consistently, among surviving chickens, 75% Cobb chickens showed extensive, score-6 lesions in the small intestinal tract, and 9% Ghs6 chickens exhibited severe score-6 lesions, while none of M5.1 chickens presented any abnormalities in the small intestine at 3 dpi (Fig. 1C). Additionally, NE significantly reduced weight gain in Cobb and Ghs6 chickens but had no impact on M5.1 chickens (Fig. 1D). These results clearly indicated that, among three breeds of chickens, M5.1 chickens are the most resistant to NE and Cobb chickens are the most susceptible, with Ghs6 being intermediate.

Differences in the ileal microbiota among three chicken breeds under healthy and NE conditions

To investigate the microbiota differences among M5.1, Ghs6, and Cobb chickens and their microbiota shifts in response to NE, bacterial DNA was isolated from the ileal and cecal digesta of mock- and NE-infected chickens and subjected to 16S rRNA gene sequencing. After quality control, 7,750,326 high-quality sequencing reads were obtained, averaging 61,510 ± 7,021 sequences per sample. Following denoising and removal of amplicon sequence variants (ASVs) present in less than 5% of samples, 399 and 421 ASVs were identified in the ileum and cecum, respectively.

Analysis of α-diversity of the ileal microbiota, as measured by Shannon Index, revealed a progressive decline across the three chicken breeds in relation to increasing resistance to NE (Fig. 2A). This trend was further accentuated following NE challenge, with the NE-resistant M5.1 chickens exhibiting the least perturbation. Although β-diversity showed significant differences among the three breeds based on weighted UniFrac distance, M5.1 chickens were the least affected by NE (Fig. 2B). The ileal microbiota profiles were drastically different among the breeds (Fig. 2C). In healthy Cobb chickens, three dominant taxa included group A Lactobacillus F1 (25.5%), Ligilactobacillus salivarius F3 (14.0%), and Corynebacterium F45 (10.3%). In contrast, Staphylococcus gallinarum (F8) was highly abundant in healthy Ghs6 (43.9%) and M5.1 chickens (36.9%) but was largely absent (0.8%) in Cobb chickens (Fig. S1A). Corynebacterium was almost undetectable in both M5.1 and Ghs6 chickens. Notably, Lactobacillus johnsonii (F4) was minimal in Ghs6 chickens (0.3%), and group A Lactobacillus accounted for only 3.9% of the total ileum bacteria. In M5.1 chickens, L. johnsonii represented 34.7%, while group A Lactobacillus comprised just 2.2% in the ileum.

Among the 40 most abundant ileal ASVs, the majority were significantly different among the three chicken breeds under healthy and NE-challenged conditions. The ileal microbiota profiles of M5.1 and Ghs6 chickens resembled each other more than that of Cobb chickens (Fig. 2D). Among three major pathobionts, C. perfringens increased drastically in NE-susceptible Cobb chickens from 0.03% to 34.1% but remained minimal in NE-challenged Ghs6 (1.5%) and M5.1 (0.08%) chickens (Fig. 2D and Fig. S1). Escherichia (F7) also significantly bloomed in NE-infected Cobb chickens but not in M5.1 or Ghs6 chickens. Enterococcus cecorum (F30) showed a significant increase in all three breeds following NE challenge. Major lactic acid bacteria (LAB) such as group A Lactobacillus and L. salivarius were lowly abundant in healthy M5.1 and Ghs6 chickens compared to Cobb chickens but were markedly enriched in both breeds following NE challenge (Fig. S1). In response to NE, L. johnsonii was significantly reduced in Cobb chickens but increased substantially in both M5.1 and Ghs6 chickens.

Differences in the cecal microbiota among three chickens breeds under healthy and NE conditions

Consistent with observations in the ileum, α-diversity of the cecal microbiota progressively declined with increasing NE resistance across the three chicken breeds, and NE infection further reduced Shannon Index in Cobb but not Ghs6 or M5.1 chickens (Fig. 3A). Moreover, β-diversity analysis using weighted UniFrac distance revealed distinct microbiota structures among the three breeds with and without NE challenge (Fig. 3B). The taxonomic composition of the cecal microbiota also varied markedly among breeds (Fig. 3C). Notably, Bacteroides fragilis (F2) was the dominant taxon in healthy Ghs6 (26.4%) and M5.1 (35.0%) chickens, but accounted for only 6.2% in healthy Cobb chickens (Fig. 3C and Fig. S2). Following NE challenge, B. fragilis members (F2 and F16) were enriched in Cobb chickens but showed a slight reduction in Ghs6 and M5.1 chickens (Fig. 3D).

Among major LAB species including Group A Lactobacillus (F1), L. salivarius (F3), and L. johnsonii (F4) were less abundant in M5.1 chickens but increased substantially following NE infection. In contrast, L. johnsonii was significantly suppressed in NE-challenged Cobb chickens. Short-chain fatty acid (SCFA)-producing bacteria such as two Faecalibacterium members (F14 and F19) were more prevalent in Cobb chickens under healthy conditions but diminished across all breeds after NE infection. Conversely, several other SCFA-producing members of the Oscillospiraceae family (e.g., F24, F50, and F59) were more abundant in M5.1 and Ghs6 chickens and declined following NE challenge, while showing enrichment in NE-infected Cobb chickens.

Regarding the three major pathobionts, C. perfringens remained at low abundance in the cecum across all breeds under both healthy and NE challenge conditions, although a statistically significant increase was observed in NE-infected Cobb and M5.1 chickens (Fig. 3D and Fig. S2). Escherichia was enriched in NE-infected Cobb chickens but remained largely unchanged in Ghs6 and M5.1 chickens. E. cecorum tended to increase in Cobb and M5.1 chickens following NE challenge, with no notable changes in Ghs6 chickens. Interestingly, two other Clostridium species (F51 and F82) showed no major alterations in response to NE across all three breeds.

Differential protection of Cobb chickens from NE by the cecal microbiota of three chicken breeds

To directly evaluate the efficacy of intestinal microbiota in alleviating NE, cecal microbiota was prepared from all three chicken breeds and transplanted to naïve Cobb chickens, followed by NE challenge (Fig. 4A). Our results showed that the M5.1 microbiota provided the best protection against NE, with 100% survival in the transplanted group, while approximately 40% of the chickens in the mock-transplanted control group died from severe intestinal lesions at 3 dpi (Fig. 4B). Interestingly, the cecal microbiota from the two susceptible breeds, Ghs.6 and Cobb, also conferred significant protection to naïve Cobb chickens, although with reduced efficacy compared to the M5.1 microbiota (Fig. 4B and 4C). Consistently, the cecal microbiota of all three chicken breeds partially reversed NE-induced weight loss in recipient Cobb chickens (Fig. 4D).

Further examination revealed obvious changes in the ileal microbiota of recipient chickens following cecal microbiota transplantation (CMT). Except for microbiota richness (Fig. S3A), CMT of the M5.1 microbiota, but not other microbiota, significantly increased the Shannon Index of the ileal microbiota of recipient chickens (Fig. S3B) and caused significant shifts in microbiota structure based on weighted UniFrac distance (Fig. S3C). The ileal microbiota composition also experienced notable alterations (Fig. S3D). CMT of all three breeds significantly enriched two Gemmiger species (F22 and F31) with a tendency to increase many but not all LAB species. In contrast, Rombousia (F23) was significantly reduced following CMT of all three breeds. Among three major pathobionts, C. perfringens and E. cecorum were unaltered, but Escherichia was significantly diminished following CMT. In response to NE, C. perfringens and Escherichia drastically bloomed in Cobb chickens mock-transplanted or transplanted with the Ghs6 or Cobb microbiota. In contrast, CMT with the M5.1 microbiota caused no blooming of C. perfringens and a significant reduction in Escherichia and E. cecorum in the ileum of recipient chickens.

CMT also caused significant changes to the cecum of recipient Cobb chickens under healthy and NE conditions (Fig. S4A-S4D). Similar to the ileum, two Gemmiger species (F22 and F31) were significantly enriched by all three transplanted microbiotas in both healthy and NE-infected recipients (Fig. S4E). Many LAB species were largely unaffected by CMT, except that group A lactobacillus and Limosilactobacillus oris (F17) were significantly enriched in recipient chickens in response to NE, with M5.1 microbiota-transplanted chickens showing the largest increase. Neither C. perfringens nor Escherichia bloomed in Cobb chickens receiving the M5.1 microbiota. Surprisingly, Bacteroides, the dominant species in the cecum of both M5.1 and Ghs6 chickens, failed to be detected in the cecum of recipient Cobb chickens following CMT, with or without NE challenge.

Protection of chickens from NE by B. pseudolongum

Both the ileal and cecal microbiotas of M5.1 or Ghs6 chickens resembled each other more than Cobb chickens under both mock- and NE-infected conditions. To explain the obvious difference in NE resistance between M5.1 and Ghs6 chickens, we detected a notable difference in the differential abundance of two Bifidobacterium species, namely B. anseris (F84) and B. pseudolongum (F201) in both the ileum and the cecum (Fig. 5A-5D) among the three chicken breeds. Both Bifidobacterium species were largely absent in both mock- and NE-infected Ghs6 and Cobb chickens as well as in mock-infected M5.1 chickens, but showed a significant enrichment in both the ileum and cecum of M5.1 chickens following NE challenge.

To directly verify if Bifidobacterium plays a role in NE resistance, we plated the cecal bacteria of M5.1 chickens on MRS plates and identified an isolate to be B. pseudolongum through Sanger sequencing of its full-length 16S rRNA gene. It showed a potent activity in inhibiting C. perfringens growth with a 5.6-fold reduction when incubated 1:1 for 24 h in a coculture assay (Fig. 5E). To further evaluate its efficacy against NE, we orally inoculated approximately 1 × 10⁷ CFU of B. pseudolongum to each Cobb chicken on days 9, 11, 13, and 15, and challenged them with E. maxima and C. perfringens to induce NE on days 10 and 14, respectively (Fig. 5F). Only 42.2% of NE-challenged chickens survived at 3 dpi without intervention, compared to 83.3% survival in those that received B. pseudolongum (Fig. 5G), which also significantly alleviated intestinal lesions (Fig. 5H), but with no obvious impact on growth (Fig. 5I). Overall, these results suggested a protective role of B. pseudolongum against NE.

DISCUSSION

Fayoumi chickens, originating from Egypt, have been found to be highly resistant to multiple diseases ^5–10. We hypothesized that, shaped by its unique genetics, this chicken breed may harbor distinct intestinal microbiota to confer enhanced disease resistance. Our results demonstrated that, among three chicken breeds studied including Fayoumi, Leghorn, and Cobb, Fayoumi chickens exhibit the highest resistance to NE, whereas Cobb chickens are the most susceptible. We further demonstrated that cecal microbiota transplantation from Fayoumi chickens offers superior protection of NE-susceptible Cobb chickens from NE. Additionally, oral administration of Bifidobacterium, a uniquely enriched commensal in NE-infected Fayoumi chickens provided significant protection of Cobb chickens against NE, suggesting the utility of intestinal bacteria and Bifidobacterium in particular in mitigating NE and perhaps other diseases.

Differential intestinal microbiota responses to NE among Cobb, Ghs6, and M5.1 chickens

The intestinal microbiome significantly influences health and disease, with its composition shaped by various factors, including host genetics ^11–13. In this study, we observed significant differences in the intestinal microbiota among three chicken breeds. For example, Bacteroides was the most abundant in the cecum of M5.1 and Ghs6 chickens but was minimally present in Cobb chickens, which is consistent with several recent analyses showing the prevalence of Bacteroides in many indigenous breeds but not in Cobb chickens ^13–16. Bacteroides is a genus of non-spore-forming, Gram-negative bacteria that degrade nondigestible carbohydrates to produce SCFAs, offering various host benefits and providing CR against pathogens like Clostridioides difficile ^17,18. However, certain Bacteroides species are opportunistic pathogens that can promote chronic inflammation ¹⁹. While Bacteroides may aid in NE resistance, its exact role in CR against NE requires further investigation. It is noteworthy that Bacteroides seems dispensable for NE resistance, as Cobb chickens become highly resistant to NE infection despite having undetectable levels of Bacteroides following M5.1 microbiota transplantation.

To our surprise, Staphylococcus, mainly S. gallinarum, was most prevalent in M5.1 and Ghs6 chickens, accounting for 35–45% of the total ileal bacteria, but was largely absent in Cobb chickens. S. gallinarum is a non-pathogenic, coagulase-negative bacterium commonly found in healthy chickens, pheasants, and humans ^20–22. It has probiotic properties, showing activity against pathogenic Escherichia coli and Klebsiella pneumoniae in vitro ²³. Additionally, it produces Staphyloferrin A, a siderophore that suppresses pathogenic bacteria growth by chelating iron, essential for virulence and bacterial interactions ²³. The role of Staphylococcus in NE resistance warrants further investigation.

Notably, NE-resistant M5.1 chickens had a significantly higher abundance of Weissella in the ileum compared to susceptible Ghs6 and Cobb chickens. Weissella, part of the Leuconostocaceae family, is known for its probiotic and anti-inflammatory potential ²⁴. For example, W. cibaria can inhibit pathogenic microorganisms through metabolites like exopolysaccharides ²⁵. Weissella species produce bacteriocins, such as Weissellicins ^25–27. The role of Weissella in poultry health and disease remains underexplored, necessitating further studies.

Additionally, our results clearly demonstrated varying abundances of LAB species among the three chicken breeds and their distinct responses to NE. Group A Lactobacillus, including highly related species like L. crispatus, L. acidophilus, and L. gallinarum ²⁸ that cannot be distinguished by the V3–V4 region of the bacterial 16S rRNA gene, were more abundant in the ileum and cecum of Cobb chickens than in M5.1 and Ghs6 chickens. However, Group A Lactobacillus remained largely unchanged by NE in Cobb chickens but was significantly enriched in M5.1 and Ghs6 chickens. Conversely, L. johnsonii was significantly reduced in Cobb chickens but enriched in M5.1 and Ghs6 chickens in response to NE. The differential response of the same LAB species in different breeds suggests the possible presence of different LAB strains, explaining the variation in the NE resistance pattern. It is important to confirm and isolate LAB strains preferentially growing in Fayoumi chickens and investigate their efficacy in disease resistance. Additionally, relative contributions of different LAB species to NE resistance require further investigation, although many LAB species have shown benefits against NE ^3,29.

Protection of naive Cobb chickens from NE through CMT

Given recent successes of transplanting fecal or cecal microbiota in conferring CR in chickens against pathogens such as Salmonella ^30,31, Campylobacter jejuni ^32–34, and C. perfringens infections ³⁵, we compared the efficacy of the cecal microbiota from three chicken breeds in providing CR against NE. We observed a drastic improvement in NE resistance among naive Cobb chickens receiving the Fayoumi microbiota. Additionally, the cecal microbiota of NE-susceptible Cobb and Ghs6 chickens also provided notable, albeit less pronounced, protection against NE. These findings align with a previous report showing reduced chicken intestinal lesions in a subclinical NE model following transplantation of bioreactor-propagated cecal microbiota of adult chickens ³⁵. However, our results contrast with an earlier study demonstrating that CMT from a resistant chicken line (ADOL Leghorn Line 6₁) to a susceptible line (ADOL Leghorn Line N) failed to confer CR against C. jejuni infection ³².

The discrepancy among these studies may be attributed to differences in the CMT preparation method. In our study and that of Zaytsoff, et al. ³⁵, the microbiota transplants were prepared under anaerobic conditions, whereas Chintoan-Uta et al. ³² prepared CMT aerobically. It is plausible that anaerobic commensal bacteria, which are crucial for disease resistance, may not survive well during aerobic microbiota preparation. Supporting this hypothesis, transplantation of anaerobic cecal microbiota was shown to provide CR against C. jejuni ³⁴. However, transplantation of both aerobically and anaerobically cultured mouse fecal microbiota offered CR against C. jejuni in chickens ³³. Further research is warranted to elucidate the specific microbial communities and mechanisms underlying the protection against different pathogens.

We observed enrichment of Gemmiger in both the ileum and cecum of recipient chickens following CMT from all three chicken breeds, whereas Megamonas and Bacteroides were enriched following transplantation of bioreactor-propagated cecal microbiota ³⁵. The differences in outcomes between the two studies likely stem from variations in the transplanted microbiota and genetic differences in recipient chickens. Gemmiger, a genus of bacteria in the family Oscillospiraceae, is closely related to Subdoligranulum and Faecalibacterium ^36,37, both of which produce SCFAs with anti-inflammatory properties. Gemmiger was reported to be depleted in multiple cohorts of inflammatory bowel disease patients, alongside other butyrate producers such as Faecalibacterium ³⁸. Administering Gemmiger or its related Faecalibacterium or Subdoligranulum may prove beneficial against NE.

Bifidobacterium -mediated protection of chickens from NE

Despite the similarity in intestinal microbiota between Ghs6 and M5.1 chickens, both of which were hatched in the same location, Ghs6 chickens are evidently more susceptible to NE than M5.1 chickens. A notable observation is the marked increase in Bifidobacterium in NE-challenged M5.1 chickens, a response not observed in Ghs6 or Cobb chickens. Oral administration of Bifidobacterium pseudolongum conferred substantial protection against NE, underscoring its protective role. This is consistent with the well-documented antibacterial and immunomodulatory properties of Bifidobacterium species ^39,40. Additionally, Bifidobacteria synthesize essential vitamins such as riboflavin, thiamine, vitamin B6, and vitamin K, along with bioactive molecules like folic acid, niacin, and pyridoxine ⁴¹. Unlike Lactobacillus species that produce both D(-)-lactic acid and L(+)-lactic acid, Bifidobacteria predominantly produce L(+)-lactic acid, which is more readily metabolized by humans and animals ⁴¹.

Additionally, Bifidobacterium has demonstrated efficacy against subclinical NE ⁴² and C. perfringens in co-culture studies ⁴³, consistent with our in vitro and in vivo observations. However, it is noted that, although Bifidobaceterium is enriched NE-challenged Fayoumi chickens and beneficial against NE, it is unlikely to be solely responsible for NE resistance in Fayoumi chickens. This is evidenced by the absence of Bifidobacterium in recipient Cobb chickens following CMT from any chicken breed, despite the robust protection observed. Therefore, it is plausible that multiple bacterial species in the intestinal microbiota act synergistically to provide CR against NE.

CONCLUSIONS

Our study demonstrates that Fayoumi chickens exhibit greater resistance to NE compared to Leghorn layers and Cobb broilers. Additionally, we provide compelling evidence that the intestinal microbiota from Fayoumi chickens confers significant protection against NE in newly-hatched Cobb chickens, with B. pseudolongum playing a crucial role in this protective effect. These findings underscore the potential of leveraging disease-resistant chicken microbiota for NE mitigation. Future research should focus on identifying the specific bacterial consortia responsible for CR and exploring their application in developing probiotic treatments to enhance poultry health and productivity.

METHODS

Ethics statement

All animal experiments described in this study were conducted according to the recommendations in the Guide for the Care and Use of Agricultural Animals in Research and Teaching, 4th edition (2020) and approved by the Institutional Animal Care and Use Committee of Oklahoma State University under protocol number AG-23-35.

NE challenge of inbred chickens

To investigate the difference of intestinal microbiota in response to NE, 48 day-of-hatch M5.1 and Ghs6 chicks, with 24 birds/breed, were obtained from Iowa State University (Ames, Iowa), while 24 day-of-hatch Cobb-500 chicks were obtained from Cobb-Vantress (Siloam Springs, Arkansas). Chickens were housed in floor pens (3' × 3') with 12 birds/pen and fresh wood shavings in an environmentally controlled room under standard management. Chickens had free access to tap water and an unmedicated mash corn-soybean starter diet containing 21.5% crude protein that meets or exceeds the nutrient requirements of the NRC recommendations ⁴⁴ throughout the study. Within each breed, animals were weighed individually on day 16 and assigned randomly to either the mock or NE group. Each animal in the NE group was orally inoculated with 1 × 10⁴ sporulated oocysts of the E. maxima M6 strain (kindly provided by Dr. John R. Barta, University of Guelph, Canada) in 1 mL PBS on day 16, followed by four sequential inoculations with approximately 5 × 10⁸ CFU of netB- and tpeL-positive C. perfringens Brenda B strain (kindly provided by Dr. Lisa Bielke, North Carolina State University, Raleigh, North Carolina) in 2 mL fluid thioglycollate (FTG) broth (Thermo Fisher Scientific) twice daily on days 20 and 21, respectively, as previously described ^45–47. The mock-infected group received 1 mL PBS or 2 mL FTG each time.

To minimize cross-contamination, floor pens were separated from each other with plastic sheets. Animals were observed twice daily for survival and behavior till day 23. Chickens reluctant to move were euthanized to minimize undue suffering. On day 23, all surviving chickens were weighed individually and sacrificed via CO₂ asphyxiation. Lesions in the small intestine were scored on a scale of 0–6 as described ⁴⁸. Additionally, the digesta in the proximal ileum (approximately 0.5 g) and cecum (approximately 0.2 g) were separately collected and stored at − 80°C for microbial genomic DNA extraction.

Preparation of the cecal microbiota

Cecal microbiota was collected from 35-day-old, healthy M5.1, Ghs6, and Cobb chickens with three birds/breed. After euthanasia via CO₂ asphyxiation, the cecum was ligated at the ileal-cecal junction, excised, and transferred into BACTRON300™ Anaerobic Chamber (Sheldon Manufacturing, Cornelius, Oregon) within 1 h. The cecal digesta was collected, combined within each breed, weighed, and diluted with five volumes (w/v) of reduced PBS. After filtration through a 70-µM cell strainer, each cecal microbiota suspension was further diluted 10-fold in PBS containing 10% glycerol and stored at − 80°C until further use. On the day of CMT, frozen microbiota suspensions were thawed at 37°C and dispensed anaerobically into 1-mL syringes attached to a feeding needle and transferred to the animal facility in resealable Ziploc® plastic bags.

Cecal microbiota transplantation

A total of 120 day-of-hatch Cobb chickens were obtained from Cobb-Vantress and randomly assigned to one of eight groups with 15 birds/group. Each animal received 0.2 mL PBS or 0.2 mL diluted cecal microbiota from healthy M5.1, Ghs6, or Cobb chickens on days 0, 1, 9, and 13. On days 10 and 14, four groups of animals were challenged with 1 × 10⁴ sporulated oocysts of E. maxima B6 and approximately 5 × 10⁸ CFU of C. perfringens Branda B to induce NE, while the other four groups were mock-infected. Animals were observed twice daily for mortalities till day 17. Chickens were weighed individually on days 0, 10, and 17. On day 17, all surviving animals were sacrificed and examined for small intestinal lesion scores. Additionally, the digesta in the proximal ileum and cecum were collected and stored at − 80°C for microbial genomic DNA extraction.

Isolation, culture, and oral administration of B. pseudolongum against NE

The ileal and cecal digesta of three 35-day-old M5.1 chickens were collected, diluted 10-fold in in reduced PBS, and filtered through a 70-µM cell strainer in an anaerobic chamber. After 10-fold serial dilutions in reduced PBS, 100 µL of each dilution was plated on de Man, Rogosa, and Sharpe (MRS) and reinforced clostridial medium (RCM) agar plates, respectively. After 24-h anaerobic culture, colony-PCR was performed with well-isolated colonies using primers (27F: AGA GTT TGA TCC TGG CTC AG and 1492R: GGT TAC CTT GTT ACG ACT T) to amplify the entire 16S rRNA gene, followed by Sanger sequencing. An isolate was identified to share 99.5% identity to B. pseudolongum and restreaked on MRS plates three times, followed by anaerobic propagation in MRS or RCM.

To evaluate the protective efficacy of B. pseudolongum against NE, 90 day-of-hatch Cobb chickens were randomly assigned to one of three treatments with 15 birds/pen and two pens/treatment. Each animal received approximately 1 × 10⁷ CFU of B. pseudolongum in 1 mL reduced PBS on days 9, 11, 13, and 15. On days 10 and 14, two groups of animals were challenged with 5 × 10³ sporulated oocysts of E. maxima B6 and approximately 5 × 10⁸ CFU of C. perfringens Branda B to induce NE, while the third group was mock-infected with PBS and FTG on respective days. Animals were observed twice daily for mortalities till day 17. Chickens were weighed individually on days 0, 10, and 17. On day 17, all surviving animals were sacrificed and examined for small intestinal lesion scores.

Bacterial coculture assay

To directly evaluate the anti-C. perfringens activity of B. pseudolongum, both bacteria were grown anaerobically in Brain Heart Infusion (BHI) broth overnight and diluted to 2 × 10⁷ CFU/mL in BHI, mixed 1:1, and incubated anaerobically for 24 h at 37°C. The survival of C. perfringens was assessed through serial plating on perfringens-selective tryptose sulfite cycloserine (TSC) agar plates (Sigma Aldrich, St. Louis, MO).

Bacterial DNA isolation and 16S rRNA sequencing

Fecal DNA MicroPrep and MiniPrep Kits (Zymo Research Irvine, CA) were used for isolation of DNA from the ileal and cecal digesta in animal trials, respectively. The concentration and quality of DNA was measured by Nanodrop One Spectrophotometer (Thermo Fisher Scientific). High-quality DNA samples were shipped on dry ice to Novogene (Beijing, China) for PE250 deep sequencing of the V3-V4 region of bacterial 16S rRNA gene using primers (341F: CCT AYG GGR BGC ASC AG and 806R: GGA CTA CNN GGG TAT CTA AT) on the Illumina NovaSeq 6000 system. PCR amplification and library preparation were performed by Novogene (Beijing, China) using NEBNext® Ultra™ Library Prep Kit (New England Biolabs, Ipswich, MA, USA), generating a minimum of 30,000 raw sequencing reads per sample.

Bioinformatics and statistical analysis

Bioinformatic analysis was conducted as we previously described ^10,49–51. Briefly, raw sequencing reads were analyzed using QIIME 2 v2023.7 ⁵². After filtration of low-quality reads, clean sequencing reads were trimmed to 402 nucleotides and denoised using Deblur ⁵³. The resulting sequences were then classified into bacterial ASVs using the RDP 16S rRNA training set (v. 18) and Bayesian classifier. A bootstrap confidence of 80% was used for taxonomic classification. ASVs with a classification confidence below 80% were assigned to the last confidently classified taxonomic level, followed by “_unclassified”. ASVs present in fewer than 5% of samples were removed from downstream analysis. The top 100 ASVs, along with all differentially enriched taxa, were further validated and reclassified, if necessary, using an updated EzBioCloud 16S database (v2023.08.23) ⁵⁴. Species-level classification was assigned to sequences sharing greater than 97% identity.

Analysis and visualization of α- and β-diversities of the microbiota composition were analyzed using the ‘phyloseq’ R package v1.46.0 ⁵⁵. To visualize the overall biodiversity and complexity within samples, the number of ASVs, Pielou’s evenness index, and Shannon index were used to calculate and display the richness, evenness, and overall diversity. The β-diversity was determined using weighted and unweighted UniFrac distances. Statistical significance in α-diversity and relative abundance for each sampling day was determined using non-parametric Mann-Whitney U test. Significance in β-diversity was determined using non-parametric permutational multivariate analysis of variance (PERMANOVA) with 999 permutations using the vegan package v. 2.6.4 ⁵⁶. P < 0.05 was considered statistically significant. Differential abundance of bacteria among different groups of chickens was determined using ANCOM-BC2 ⁵⁷.

Abbreviations

ASV Amplicon sequence variant

BHI Brain heart infusion broth

CMT Cecal microbiota transplantation

CR Colonization resistance

dpi Days post-infection

LAB Lactic acid bacteria

MRS de Man, Rogosa, and Sharpe broth

NE Necrotic enteritis

RCM Reinforced clostridial medium

SCFA Short-chain fatty acid

Data Availability

Raw sequencing reads of this study was deposited in the NCBI GenBank SRA database under the accession number PRJNA1132701.

Acknowledgement

We would like to thank Dr. John R. Barta at the University of Guelph, Canada for kindly providing E. maxima strain M6. We are grateful to Dr. Lisa Bielke at North Carolina State University for providing the C. perfringens strain Brenda B. We also thank Ms. Zijun Zhao for helping with animal handling.

Author Contribution

JL, JG, IT, MAW, PP, AS, and GZ conducted animal trials; JL and JG processed the samples; JL, JG, IT, and GZ analyzed the data; JL drafted the manuscript; GZ, MGK, and SJL revised the manuscript; GZ conceived and supervised the study. All authors reviewed the manuscript and agreed to the published version of the manuscript.

Competing interests

The authors declare no competing interests.

Supplementary Information

Fig. S1

Relative abundances (%) of selected ileal bacteria among three chicken breeds under healthy and NE conditions. Fig. S2. Relative abundances (%) of selected cecal bacteria among three chicken breeds under healthy and NE conditions. Fig. S3. Alternations of the ileal microbiota of Cobb chickens following cecal microbiota transplantation (CMT) from three chicken breeds. Fig. S4. Alternations of the cecal microbiota of Cobb chickens following cecal microbiota transplantation (CMT) from three chicken breeds.

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Fig. 1

Differential susceptibility of three chicken breeds to necrotic enteritis (NE). (A) Experimental scheme. Three chicken breeds including Fayoumi M5.1, Leghorn Ghs6, and Cobb chickens were randomly assigned to the mock or NE group with 12 birds/group. Chickens in the NE group were subjected to an initial challenge with Eimeria maxima on d 16 and four subsequent challenges with Clostridium perfringens twice daily on d 20 and 21, while the remaining chickens were mock-infected. (B) Animal survival rate (%) between days 20–23. *P < 0.05 and ***P < 0.001 compared to NE-challenged Cobb chickens, based on the log-rank test. (C) The frequency (%) of small intestinal lesion scores (LS) of surviving chickens on d 23. *P < 0.05 and ***P < 0.001 based on the Kruskal-Wallis test and post-hoc Dunn’s test. (D) Average individual body weight gains (g) of surviving animals between d 16–23. Data shown are mean ± SEM. Means not sharing a common superscript letter denote statistical significance (P < 0.05) based on one-way ANOVA and post-hoc Tukey’s test.

Fig. 2

Differences in the ileal microbiota among three chicken breeds under healthy and NE conditions. M5.1, Ghs.6, and Cobb chickens were either mock-infected or subjected to NE challenge (n = 12 per group). Ileal digesta samples were collected from surviving animals on d 23 for bacterial DNA isolation and 16S rRNA gene sequencing. (A) Box and whisker plot depicting Shannon Index across different treatment groups. Significance was assessed using the Kruskal–Wallis test and post-hoc Dunn’s test. Different superscript letters denote significance (P < 0.05) in pairwise comparisons. (B) Principal coordinates analysis (PCoA) plot of weighted UniFrac distances. Significance was determined using PERMANOVA. (C) Relative abundances (%) of the top 15 ASVs in the ileal microbiota. (D) Heatmap showing NE-induced differential enrichment of the top 40 ASVs in the ileum. The bottom panel depicts log2 fold changes (log2FC) in ileal bacterial abundance across six groups relative to mock-infected Cobb chickens. Groups not sharing a common superscript letter in a column denote statistically significant differences (P < 0.05) based on ANCOM-BC2 analysis ⁵⁷. The top panel indicates log2FC values comparing NE-infected chickens to their respective mock-infected controls. *P-adjusted < 0.05, **P-adjusted < 0.01, and ***P-adjusted < 0.001 as determined by the Kruskal-Wallis test and post-hoc Dunn’s test with Benjamini-Hochberg correction.

Fig. 3

Differences in the cecal microbiota among three chicken breeds under healthy and NE conditions. M5.1, Ghs.6, and Cobb chickens were either mock-infected or subjected to NE challenge (n = 12 per group). Cecal digesta samples were collected from surviving animals on d 23 for bacterial DNA isolation and 16S rRNA gene sequencing. (A) Box and whisker plot depicting Shannon Index across different treatment groups. Significance was assessed using the Kruskal–Wallis test and post-hoc Dunn’s test. Different superscript letters denote significance (P < 0.05) in pairwise comparisons. (B) Principal coordinates analysis (PCoA) plot of weighted UniFrac distances. Significance was determined using PERMANOVA. (C) Relative abundances (%) of the top 15 ASVs in the cecal microbiota. (D) Heatmap showing NE-induced differential enrichment of the top 50 ASVs in the cecum. The bottom panel depicts log2 fold changes (log2FC) in bacterial abundance across six groups relative to mock-infected Cobb chickens. Groups not sharing a common superscript letter in a column denote statistically significant differences (P < 0.05) based on ANCOM-BC2 analysis ⁵⁷. The top panel indicates log2FC values comparing NE-infected chickens to their respective mock-infected controls. *P-adjusted < 0.05, **P-adjusted < 0.01, and ***P-adjusted < 0.001 as determined by the Kruskal-Wallis test and post-hoc Dunn’s test with Benjamini-Hochberg correction.

Fig. 4

Differential protection of Cobb chickens from NE by the cecal microbiota of three chicken breeds. (A) Experimental scheme. A total of 120 day-of-hatch Cobb chicks were randomly divided into eight groups (n = 15 per group), with each receiving the cecal microbiota prepared from M5.1, Ghs6, or Cobb chickens or an equal volume of PBS via oral gavage on days 0, 1, 9, and 13. Four groups were challenged with E. maxima on day 10 and C. perfringens on day 14 to induce NE, while the other four groups receiving the same volume of PBS or fluid thioglycollate broth (FTG) on respective days. (B) Animal survival (%) between day 14–17 among four groups of NE-infected Cobb chickens that received PBS or the cecal microbiota from Cobb, Ghs6, or M5.1 chickens. *P < 0.05 and ***P < 0.001 compared to the NE group receiving only PBS, based on the log-rank test. (C) The frequency (%) of intestinal lesion scores (LS) among four groups of NE-infected Cobb chickens that received PBS or the cecal microbiota on day 17. **P < 0.001 based on the Kruskal-Wallis test and post-hoc Dunn’s test. No other groups are significantly different. (D) Average body weight gains of surviving animals among eight groups of recipient Cobb chickens between day 10–17. Data shown are means ± SEM. Means not sharing a common superscript letter denote statistical significance (P < 0.05) based on one-way ANOVA and post-hoc Tukey’s test.

Fig. 5

Protection of chickens from NE by Bifidobacterium pseudolongum. Relative abundances (%) of B. anseris (A and C) and B. pseudolongum (B and D) in the ileum and cecum of healthy and NE-challenged M5.1, Ghs6, and Cobb chickens (n = 12 per group). (E) C. perfringens counts following 24-h incubation with or without B. pseudolongum at the 1:1 ratio. (F) Experimental scheme assessing B. pseudolongum-mediated protection against NE. A total of 90 day-of-hatch Cobb chicks were randomly divided into three groups (n = 30 per group), with each receiving B. pseudolongum or an equal volume of PBS via oral gavage on days 9, 11, 13, and 15. Two groups were challenged with E. maxima on day 10 and C. perfringens on day 14 to induce NE, while the third group receiving the same volume of PBS or fluid thioglycollate broth (FTG) on respective days. (G) Animal survival (%) between day 14–17 among three groups. **P < 0.01 compared to the NE group receiving only PBS, based on the log-rank test. (H) The frequency (%) of intestinal lesion scores (LS) among three groups. **P < 0.001 based on the Kruskal-Wallis test and post-hoc Dunn’s test. (I) Average body weight gains of animals among three groups between day 10–17. Data shown are means ± SEM. Means not sharing a common superscript letter denote statistical significance (P < 0.05) based on one-way ANOVA and post-hoc Tukey’s test.

Supplementary Information

Fig. S1

Relative abundances (%) of selected ileal bacteria among three chicken breeds under healthy and NE conditions. M5.1, Ghs.6, and Cobb chickens were either mock-infected or subjected to NE challenge (n = 12 per group). Ileal digesta samples were collected from surviving animals on d 23 for bacterial DNA isolation and 16S rRNA gene sequencing. Relative abundances (%) of selected bacteria across six groups are shown. Groups not sharing a common superscript letter denote statistically significant differences (P < 0.05) based on ANCOM-BC2 analysis ⁵⁷.

Fig. S2

Relative abundances (%) of selected cecal bacteria among three chicken breeds under healthy and NE conditions. M5.1, Ghs.6, and Cobb chickens were either mock-infected or subjected to NE challenge (n = 12 per group). Cecal digesta samples were collected from surviving animals on d 23 for bacterial DNA isolation and 16S rRNA gene sequencing. Relative abundances (%) of selected bacteria across six groups are shown. Groups not sharing a common superscript letter denote statistically significant differences (P < 0.05) based on ANCOM-BC2 analysis ⁵⁷.

Fig. S3

Alternations of the ileal microbiota of Cobb chickens following cecal microbiota transplantation (CMT) from three chicken breeds. A total of 120 day-of-hatch Cobb chicks were randomly divided into eight groups (n = 15 per group), with each receiving the cecal microbiota prepared from M5.1, Ghs6, or Cobb chickens or an equal volume of PBS via oral gavage on days 0, 1, 9, and 13. Four groups were challenged sequentially with E. maxima and C. perfringens to induce NE, with the other four groups being mock-infected. The ileal digesta was collected from surviving animals, followed by DNA isolation and 16S rRNA gene sequencing. (A) Box and whisker plot of observed amplicon sequence variants (ASVs) among different groups. (B) Box and whisker plot of the Shannon Index among different groups. Significance was measured using the Kruskal–Wallis test and post-hoc Dunn’s test. Different superscripts denote significance (P < 0.05) in pairwise comparisons. (C) Principal coordinates analysis (PCoA) plots of weighted UniFrac distances. Significance was determined using PERMANOVA. (D) Relative abundances (%) of the top 15 ASVs in the ileal microbiota. (E) Heatmap showing NE-induced differential enrichment of the top 40 ASVs in the ileum. Data in the bottom two panels was depicted as log2 fold changes (log2FC) of the ileal bacteria in eight groups of chickens relative to mock-infected chickens, while the top panel indicates log2FC of the ileal bacteria in NE-infected chickens receiving the M5.1, Ghs6, or Cobb microbiota relative to NE-infected chickens receiving only PBS. *P-adjusted < 0.05, **P-adjusted < 0.01, and ***P-adjusted < 0.001 as determined by the Kruskal-Wallis test and post-hoc Dunn’s test with the Benjamini-Hochberg correction.

Fig. S4

Alternations of the cecal microbiota of Cobb chickens following cecal microbiota transplantation (CMT) from three chicken breeds. A total of 120 day-of-hatch Cobb chicks were randomly divided into eight groups (n = 15 per group), with each receiving the cecal microbiota prepared from M5.1, Ghs6, or Cobb chickens or an equal volume of PBS via oral gavage on days 0, 1, 9, and 13. Four groups were challenged sequentially with E. maxima and C. perfringens to induce NE, with the other four groups being mock-infected. The cecal digesta was collected from surviving animals, followed by DNA isolation and 16S rRNA gene sequencing. (A) Box and whisker plot of observed amplicon sequence variants (ASVs) among different groups. (B) Box and whisker plot of the Shannon Index among different groups. Significance was measured using the Kruskal–Wallis test and post-hoc Dunn’s test. Different superscripts denote significance (P < 0.05) in pairwise comparisons. (C) Principal coordinates analysis (PCoA) plots of weighted UniFrac distances. Significance was determined using PERMANOVA. (D) Relative abundances (%) of the top 15 ASVs in the cecal microbiota. (E) Heatmap showing NE-induced differential enrichment of the top 50 ASVs in the cecum. Data in the bottom two panels was depicted as log2 fold changes (log2FC) of the cecal bacteria in eight groups of chickens relative to mock-infected chickens, while the top panel indicates log2FC of the cecal bacteria in NE-infected chickens receiving the M5.1, Ghs6, or Cobb microbiota relative to NE-infected chickens receiving only PBS. *P-adjusted < 0.05, **P-adjusted < 0.01, and ***P-adjusted < 0.001 as determined by the Kruskal-Wallis test and post-hoc Dunn’s test with the Benjamini-Hochberg correction.

Yes