Early-Life Poultry-Derived Lactobacilli Drive Microbial Succession and Gut Immune Modulation in Broiler Chickens

Early-Life Poultry-Derived Lactobacilli Drive Microbial Succession and Gut Immune Modulation in Broiler Chickens | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Early-Life Poultry-Derived Lactobacilli Drive Microbial Succession and Gut Immune Modulation in Broiler Chickens Shreeya Sharma , Anna Seekatz , Mohammadali Alizadeh , Hosni Hassan , Alexander Yitabrek , Scott Pratt , Khaled Abdelaziz doi: https://doi.org/10.1101/2025.08.01.668251 Shreeya Sharma 1 Department of Animal and Veterinary Science, Clemson University , Clemson, SC 29634, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Anna Seekatz 2 Department of Biological Sciences, Clemson University , Clemson, SC 29631, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Mohammadali Alizadeh 3 Department of Pathobiology, Ontario Veterinary College, University of Guelph , Guelph, Ontario N1G 2W1, Canada Find this author on Google Scholar Find this author on PubMed Search for this author on this site Hosni Hassan 4 Prestage Department of Poultry Science, North Carolina State University , NC 27607, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Alexander Yitabrek 5 Department of Animal & Food Sciences, University of Delaware, 531 S College Ave , Newark, DE 19716, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Scott Pratt 1 Department of Animal and Veterinary Science, Clemson University , Clemson, SC 29634, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site Khaled Abdelaziz 1 Department of Animal and Veterinary Science, Clemson University , Clemson, SC 29634, USA 6 Clemson University School of Health Research (CUSHR) , Clemson, SC 29634, USA Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: Khaled2{at}clemson.edu Abstract Full Text Info/History Metrics Preview PDF Abstract Probiotic supplementation supports poultry gut health by modulating microbiome and promoting immune development, yet limited information is known about the effects of early, particularly embryonic, supplementation. In this study, we investigated the effects of administering a lactobacilli cocktail in ovo (embryonic day 18), post-hatch, or both on gut immunity and the succession of the cecal microbiota in broilers over five weeks. 16S rRNA gene-based sequencing of cecal contents revealed a steady increase in Shannon diversity during the first three weeks (PERMANOVA, p < 0.005), with community structure stabilizing by week 3 across all groups. In ovo lactobacilli administration improved early hatch rates and modulated microbial composition during early succession, including reductions in Klebsiella and Enterococcus , and enrichment of Lactobacillus , during the first two weeks (MaAsLin2, q < 0.25). These microbiome shifts were accompanied by a reduced expression of pro-inflammatory cytokines (IFN-γ, IL-1β, and IL-8) in cecal tonsils. These findings highlight the transient yet critical role of early-life probiotic interventions in shaping gut microbial colonization and immune response in broiler chickens. More importantly, a single in ovo lactobacilli dose yielded effects comparable to weekly oral or combined administration. Introduction The avian gut microbiome is a complex and dynamic ecosystem that influences digestion, absorption, immune system development, host metabolism, organ development and overall performance 1 . In modern broiler production, chicks have no direct contact with adult birds and thus acquire their initial microbiota primarily from the hatching environment. As a result, neonatal chicks are left susceptible to early colonization by opportunistic pathogens such as Escherichia coli and Salmonella spp., which can compromise gut health during the immediate post-hatch period 2 , 3 . Dietary intervention, including supplementation of probiotics, has been recognized as a strategy to support early gut health and microbiota development 4 – 6 . Probiotics are defined as “live, non-pathogenic microorganisms which, when administered in adequate amounts, confer a health benefit on the host.” 7 They exert their benefits by competitively excluding pathogens, producing antimicrobial compounds such as bacteriocins, generating organic acids like lactic acid and butyrate, which not only create an unfavorable environment for enteric pathogens but also modulate the host immune system and promote gut homeostasis 8 – 10 . As early colonizers of the poultry gut, lactic acid bacteria (LAB) are recognized for their probiotic properties 11 . They belong to the phylum Firmicutes, and species such as L. reuteri , L. acidophilus , L. crispatus and L. animalis are often isolated from healthy chicken intestines, crop or ceca 12 . Various LAB strains have been shown to confer protection against enteric pathogens such as Salmonella , E. coli , Campylobacter and Clostridium perfringens . For example, in broilers challenged with Salmonella , Lactobacillus rhamnosus GG significantly reduced pathogen loads in the ceca, liver, and spleen while modulating gut microbiota composition 13 . L. acidophilus has also been shown to enhance gut barrier integrity and reduce inflammatory cytokine expression under E. coli challenge 14 . Similarly, L. salivarius significantly reduced cecal colonization by Campylobacter , with some strains achieving over 2-log reductions in pathogen load without disrupting microbiome balance 15 . Several Lactobacillus species, including L. johnsonii , L. crispatus , L. salivarius , and L. reuteri , have been associated with decreased intestinal lesions, modulation of gut cytokine expression, and stabilization of gut microbiota in response to Clostridium perfringens infection 16 . As such, these species may serve as suitable probiotics to influence the early succession of the microbiota. While studies have evaluated the early impact of in ovo administered LAB on the microbiome of chicks 17 – 20 , their influence on weekly microbial succession throughout the rearing period has not been previously established. Colonization of the chick gut microbiome begins immediately after hatching. Initially, the microbial community is dominated by facultative anaerobes from the environment (such as Enterobacteriaceae ), but within a week, the succession shifts towards Firmicutes like Lactobacillus, with the microbiome stabilizing within the first two to three weeks of age 21 . A recent study showed that the relative abundance of Lactobacillus changed rapidly from day-old chicks (1-6%) to week-old chicks (40%) 22 . Early colonizers like Lactobacillus also engage in molecular crosstalk with the host. The gut microbiome and host immune system communicate through interactions between bacterial components and pattern recognition receptors on the intestinal epithelium and various immune cells 23 – 25 . Manipulation of the gut microbiome of hatched chicks using probiotic lactobacilli has been found to be a useful approach to modulating both gut immunity and microbiome composition 8 . However, it is widely recognized that continuous administration is typically required to achieve stable and lasting colonization 26 . This could be attributed to the fact that most of these studies supplement exogenous non-poultry-specific probiotics to the existing microbiome in hatched chicks. As a result, these probiotics may not be able to persistently colonize the intestinal tract and, therefore, exert transient protective effects, requiring continuous supplementation 27 . Early administration of probiotics through in ovo inoculation may enhance the gut microbiome composition of neonatal chicks by seeding the developing gut and competitively excluding pathogens 28 – 30 . Given the importance of early microbiome establishment and immune system maturation in newly hatched chicks 23 , 30 , 31 , this study was carried out to evaluate and compare the effects of in ovo and post-hatch administration of well-characterized, poultry-derived probiotic lactobacilli on the succession, diversity and taxonomic composition of cecal microbial communities as well as gut immune responses in broiler chicks from hatching to five weeks of age. Results In ovo administration of probiotic lactobacilli positively influenced hatchability outcomes Poultry-specific probiotic cocktail containing Lactobacillus reuteri , L. crispatus , L. animalis , and L. acidophilus (10 7 CFU/mL) was administered with a dosing volume of 100 µL following 31 . Group A (control) received no probiotics, Group B received in ovo probiotics only, Group C received both in ovo and weekly oral probiotic supplementation, and Group D received oral probiotics post-hatch only ( Figure 1A ). Chicks began hatching on embryonic day 20, with the probiotic-treated group exhibiting a higher early hatch rate (8.89%, 12/135 eggs) compared to the untreated group (4.45%, 6/135 eggs). Overall hatchability was also slightly higher in the probiotic-treated group (99.26%, 134/135 eggs) than in the untreated control group (97.04%, 131/135 eggs), reflecting a 2.2% improvement in hatchability associated with probiotic administration ( Figure 1B ). To investigate the effect of early Lactobacillus supplementation on cecal microbial succession, we performed 16S rRNA gene sequencing on cecal samples collected weekly from four experimental groups of broiler chickens (8 samples/group/week) raised under standard conditions ( Figure 1A ). Download figure Open in new tab Figure 1. Experimental design of the study and hatchability results. Microbial succession in the broiler cecum is marked by increased diversity and Clostridiales dominance The results revealed that cecal microbiota diversity increased steadily during the first three weeks (Shannon diversity, mean week 1 = 3.00), stabilizing by week three (means in weeks 2, 3, 4, 5 = 3.60, 3.87, 4.07, and 4.00) ( Figure 2A ). This increase was observed in all four treatment groups, indicating steady microbial succession independent of lactobacilli administration. Similar patterns of microbial structural changes, as assessed by nonparametric multidimensional scaling (NMDS) of the Bray-Curtis distance calculated from ASVs, were also observed over time ( Figure 2B ). No difference in overall microbiota structure was observed by the end of the experiment (weeks 3 – 5). However, comparison of the microbiota structure at each week demonstrated significant separation of untreated chickens from the remaining treatment groups at weeks 1, 2, and 3, suggesting some effect of lactobacilli manipulation on microbial succession (PERMANOVA, p < 0.005). Download figure Open in new tab Figure 2. Diversity increases early during microbial succession of the broiler chicken cecum. A) Shannon diversity index calculated from amplicon sequence variants (ASVs) in broiler chicken ceca within untreated (group A) chickens or chickens treated with lactobacilli in ovo (group B), in ovo and orally (group C), or orally (group D). Dunn’s test, * p < 0.05, ** p < 0.005, *** p < 0.0005 for comparisons within each group; # p < 0.05 for comparisons within each week across groups (week 2, A:B groups only significant finding). B) Non-metric multidimensional (NMDS) scaling of the Bray-Curtis dissimilarity index calculated from ASVs. PERMANOVA, p 2%) in each treatment group, per week. The overall phyla observed to inhabit the broiler chicken ceca were limited to Bacillota (Firmicutes, overall mean = 96.2%) and Pseudomonadata (Proteobacteria, 3.64%). At the order level, Clostridiales represented the most abundant group (overall mean = 73.64%) across all samples, followed by Lactobacillales (10.5%), Erysipelotrichales (8.85%), and Enterobacteriales (3.64%). There was a slight increase in Clostridiales in all treatment groups from week 1 (mean = 63.9%) to week 2 (80.5%), represented by increases in an unclassified Lachnospiraceae genus (week 1 mean = 22.8%, week 2 = 41.3%) and decreases in Blautia (week 1 = 16.5%, week 2 = 7.63%) from week 1 to week 2 that stabilized across subsequent weeks. In contrast, Enterobacteriales decreased from week 1 (8.0%) compared to subsequent weeks (means at weeks 2, 3, 4, and 5 = 1.79%, 1.74%, 2.41% and 2.41%). Most of this decrease over time was explained by decreased levels of Klebsiella (week 1 = 8.0%, week 5 = 0.008%) and increases in the Escherichia/Shigella genus (week 1 = 0%, week 5 = 4.26%). Lactobacilli administration influences early microbial succession Given that our PERMANOVA analysis at weeks 1 and 2 suggested divergent community structures between the treatment groups, we sought to identify compositional factors influencing these differences. We compared genus-level compositional data totaled from individual ASVs to decrease noise at each week. Using MaAsLin2 32 , we identified several differentially abundant genera across the weekly comparisons ( Figure 3A ). Many differentially abundant genera were Clostridiales, the most abundant order identified across the whole dataset; however, these genera did not necessarily demonstrate consistent patterns (i.e., consistent decreases or increases) across subsequent weeks. Several genera displayed consistent differences during weeks 1 and 2. More specifically, Klebsiella was significantly decreased in the in ovo treatment group (B) compared to the untreated, control group (A), dually treated group (C), or orally treated group (D) ( Figure 3B , Dunn’s test, p < 0.05), but decreased rapidly to almost zero in weeks 3, 4, and 5 in all groups. Enterococcus was also decreased in all treated groups compared to the control group A in weeks 1 and 2, but increased in all three treated groups in weeks 3, 4, and 5 ( Figure 3B , Dunn’s test, p < 0.05). In contrast, Lactobacillus increased in all treatment groups compared to the control in week 1 and remained elevated in most of the treatment groups throughout week 4 ( Figure 3C , Dunn’s test, p < 0.01). These results suggest that lactobacilli interventions primarily influence lactobacilli and other taxa during early microbial succession. Download figure Open in new tab Figure 3. Lactobacilli treatment influences early microbial succession in the broiler chicken cecum. A) Relative abundance (log 10 -transformed) of differentially abundant genera across treatment groups (A = control; B = in ovo , C = in ovo and oral, D = oral lactobacilli) within each week, as identified by MaAsLin2. (Linear model with BH correction, q ≤ 0.001). Relative abundance (log10-transformed) of B) Klebsiella , C) Enterococcus , and D) Lactobacillus in each treatment group by week. Dunn’s test, * p < 0.05, ** p < 0.005, *** p < 0.005. In ovo lactobacilli administration alters early gut microbial community structure To assess the impact of in ovo probiotic delivery on early gut community structure, we performed NMDS ordination based on Bray–Curtis dissimilarities. At week 1, NMDS plots revealed clear clustering among treatment groups, and PERMANOVA confirmed a significant treatment effect (F = 1.919, p = 0.001), with treatment explaining 17% of the variance in microbial composition. Furthermore, pairwise PERMANOVA comparisons between the control group (A) and the in ovo group (B) demonstrated a significant shift (F = 1.878, p = 0.001), accounting for 12% of the variation. At week 2, PERMANOVA continued to show significant differences across treatment groups (F = 1.388, p = 0.001), with group-level variance explaining 12.9%. The in ovo group (B) remained significantly different from the control (A), with 14.6% of the variance explained (F = 2.394, p = 0.001). Together, these findings indicate that a single in ovo dose of probiotics was sufficient to induce early restructuring of the cecal microbiota two weeks post-hatch ( Figure 4A ). Download figure Open in new tab Figure 4. MaAsLin2 analysis reveals early genus-level shifts driven by lactobacilli administration. A) Non-metric multidimensional scaling (NMDS) plots of Bray–Curtis dissimilarities showing cecal microbiota structure at week 1 and week 2 across treatment groups. Each point represents an individual sample. B) Barplots display genus-level relative abundances of Klebsiella , Lactobacillus , and Enterococcus across treatment groups (A: control, B: in ovo, C: in ovo + oral, D: oral only). Differential abundance testing was performed using MaAsLin2, with group A as the reference. C) Heatmap depicting differentially abundant genera across groups B ( in ovo ), C ( in ovo and oral), and D (oral), compared to the untreated control group A. Genera with positive associations are colored red (increased relative abundance), while blue indicates negative associations (reduced relative abundance), with color intensity representing the signed log-transformed FDR-adjusted p-values (–log(q-value) × sign(coefficient)). MaAsLin2 analysis reveals early genus-level shifts driven by lactobacilli administration To identify microbial taxa associated with lactobacilli administration, we applied MaAsLin2 to genus-level relative abundance data, using the control group (A) as reference. At week 1, MaAsLin2 revealed that Lactobacillus was significantly enriched in all probiotic-treated groups, with the highest increase in the in ovo group (B) (FDR = 7.996e–02; Coefficient = 1.44; Figure 4B ). In contrast, opportunistic genera Enterococcus and Klebsiella were significantly reduced in all treated groups relative to controls. Enterococcus was nearly undetectable in groups B, C, and D but prominent in group A (FDR = 2.623e–09; Coefficient = –9.70), while Klebsiella showed the greatest depletion in group B (FDR = 7.996e–02; Coefficient = –2.44). These findings suggest that early-life probiotic administration, particularly via in ovo , shows early genus-level shifts. To further resolve taxonomic drivers of community differences at week 1, we used MaAsLin2 to identify significantly associated genera. The resulting heatmap ( Figure 4C ) shows differentially abundant genera across the groups B (in ovo) , C ( in ovo + oral), and D (oral), compared to the control (A). Enterococcus and Klebsiella were significantly reduced, consistent with prior Dunn’s test analysis, while Lactobacillus , Romboutsia , and Anaerotruncus were positively associated with the lactobacilli treatment. Lactobacilli administration suppresses early expression of pro-inflammatory cytokines in cecal tonsils The expression of interferon (IFN)-γ, interleukin (IL)-1β, IL-6, and IL-8 genes in the cecal tonsils (6 cecal tonsils/group/week) was measured from the first week to the fifth week of age using RT-qPCR. During the first week, the expressions of IFN-γ, IL-1β, and IL-8 were downregulated in the oral group (P<0.05, P<0.01, and P<0.01, respectively) compared to the control. ( Figure 5 ) Similarly, in the in ovo group, IFN-γ and IL-8 were significantly reduced (P<0.05), while in the in ovo and oral combined group, IL-1β and IL-8 were significantly reduced (P<0.05). A lower, but not statistically significant, decrease in the expression of IL-6 was observed in the in ovo and oral groups compared to the control group. From weeks 2-5, there were no differences observed in the expression of any of these genes across all treatment groups (data not shown). Download figure Open in new tab Figure 5. Relative gene expression of proinflammatory cytokines (IFN-γ, IL-1β, IL-6, and IL-8) in the cecal tonsils in the first week of age. The expression of the target genes was calculated relative to the housekeeping gene (β-actin). Statistical significance among treatment groups was calculated using a non-parametric test. Error bars represent the standard error of the mean (SEM). An asterisk (*) indicates a significant difference (P<0.05), a double asterisk (**) indicates a significant difference (P<0.01) and a triple asterisk (***) indicates a significant difference (P<0.001) among the treatment groups. Discussion The chicken gut microbiome is widely recognized to influence gut development, immune function, and colonization resistance against enteric pathogens 33 . Evidence suggests that early life is important for the acquisition and establishment of the gut microbiome, during which initial microbial colonizers shape gut community structure and the maturation of the gut-associated immune system 34 . The establishment of a healthy gut microbiome during early life is thus a key determinant of intestinal development, immune maturation, and resistance to pathogen colonization in poultry 24 , 33 . Probiotic interventions have been shown to modulate cecal microbial diversity, enhance metabolic activity, and influence mucosal immune responses when administered post-hatch 35 . However, the extent to which these effects can be enhanced through early manipulation, such as in ovo probiotic administration, remains underexplored 2 , 31 , 36 , 37 . In this study, we compare in ovo , post-hatch, and combined administration of a lactobacilli cocktail to assess their effects on microbial succession, taxonomic composition, and mucosal immune responses in the chicken cecum. At the compositional level, our 16S rRNA gene-based study demonstrated that microbial diversity increased steadily over time across all groups, reflecting normal microbial succession in the broiler cecum regardless of the Lactobacillus treatment. Birds receiving lactobacilli treatments exhibited earlier and higher colonization by beneficial taxa such as Lactobacillus , Lachnospiraceae , and Clostridiales while showing significantly reduced levels of opportunistic genera like Klebsiella and Enterococcus . In contrast, the cecal microbiota of untreated birds was initially dominated by Klebsiella and Enterococcus , which declined over time as microbial diversity increased, an observation consistent with previous reports of early-life microbial succession 23 . Both Enterococcus and Klebsiella are frequently isolated from poultry environments such as feed and litter and can form persistent biofilms, exhibit biocide tolerance, and survive harsh conditions 38 – 40 . A study found out that multidrug resistant Klebsiella had high prevalence up to 35% in retail poultry meat 41 . Although some Enterococcus strains are used as probiotics, many carry antibiotic-resistant genes for tetracycline and beta-lactams 42 . Multiple studies have shown that in ovo probiotic administration leads to early enrichment of Lactobacillus and suppression of opportunistic genera in the gut 26 , 27 , 43 , 44 . For example, Wilson et al. demonstrated that chicks treated in ovo with adult microflora exhibited accelerated establishment of mature microbiota with significantly higher Lactobacillaceae/ Lactobacillus abundance and decreased abundance of Enterococcaceae/ Enterococcus at hatch compared to controls 26 . However, the study monitored microbial dynamics only up to day 10 post-hatch, at which point the observed changes had largely stabilized. In a comparable manner, Arreguin-Naca and colleagues administered a defined adult hen microbiota in ovo and observed a significant reduction in Enterococcaceae/ Enterococcus and Enterobacteriaceae at hatch, accompanied by an increase in beneficial butyrate-producing taxa such as Ruminococcus and Butryricoccus 45 . Likewise, Pedroso et al. used a complex adult-derived competitive exclusion culture and found that probiotic-treated chicks had higher microbial diversity and early colonization by beneficial taxa, aligning with our observations 46 . Furthermore, studies show that microbiota community differences between treated and untreated birds converge by approximately 3-6 weeks, reinforcing our findings 29 , 43 , 47 . We also observed that Lactobacillus itself was a differentiating genus in probiotic-treated vs control birds. Beneficial microbes such as Lactobacillus can restrict growth of these opportunistic pathogens through multiple complementary mechanisms in the gut, such as production of inhibitory metabolites, competitive exclusion, enhanced barrier function and modulation of the immune system 48 , 49 . Moreover, an increase in Lactobacillus is negatively correlated with other genera, limiting their growth likely due to niche competition 50 . Our findings coincide with previous studies that have also demonstrated the effectiveness of early lactobacilli seeding in promoting rapid gut colonization 23 , 46 , 51 , 52 . Additionally, probiotic-treated chicks showed a higher relative abundance of Clostridiales compared to controls. Members of this order, including Lachnospiraceae and Ruminococcaceae , produce butyrate and other short-chain fatty acids that support gut health, nutrient absorption, and barrier integrity 53 , 54 . They are typically dominant in mature chicken cecum, suggesting that early probiotic supplementation may accelerate microbial succession toward a more adult-like microbiota. In addition to the lactobacilli shifts in the microbiome composition, our findings demonstrated a reduction in the expression of proinflammatory cytokines, including IFN-γ, IL-1β, IL-8 and IL-6 in cecal tonsils during the first week of age. These effects may be attributed to the direct action of lactobacilli and/or to lactobacilli-induced alterations in gut microbiome composition, including an increased abundance of butyrate-producing bacteria and the resulting butyrate production, which has been reported to reduce gut inflammation. Aside from their mechanisms of action, prior research has shown that lactobacilli-mediated protection against Salmonella is linked to reduced expression of IFN-γ and IL-8 triggered by Salmonella infection 55 . It is therefore plausible to speculate that the decreased expression of these cytokines observed in the cecal tonsils of chicks given Lactobacillus either in ovo or immediately after hatching may contribute to improved resistance to Salmonella during early life. Similarly, a study has shown that dietary supplementation with Lactobacillus in broilers has been shown to markedly attenuate Salmonella -induced elevations in key inflammatory cytokines, including IL-1β, IL-8, IL-12, and IFN-γ, when compared to untreated, infected controls 56 . While not investigated in this study, previous research on fecal transplantation in chicks has reported a negative correlation between inflammatory cytokine expression and growth performance 57 . Therefore, further studies are warranted to evaluate whether in ovo lactobacilli supplementation can improve growth performance and enhance resistance to Salmonella infection in newly hatched chicks. Collectively, these results underscore the potential of a single in ovo dose of lactobacilli to modulate the intestinal microbiota and the immune system comparable to those achieved through repeated post-hatch dosing. In ovo probiotic administration allows for uniform dosing across embryos, facilitating early microbial colonization upon hatch. Previous studies have similarly demonstrated that early microbial seeding can promote mucosal immune development, enhance gut barrier function, and increase resistance to enteric pathogens during the critical first weeks of life 26 , 45 thus reducing the need for continuous supplementation of probiotics post-hatch. In conclusion, our findings indicate that in ovo Lactobacillus supplementation modulated the composition of the cecal microbiota in hatched chicks by enriching the abundance of Lactobacillus while reducing the abundance of Enterococcus and Klebsiella , underscoring the potential of early microbial interventions to shape gut microbial succession. The increased abundance of beneficial taxa alongside reduced inflammatory gene expression further supports the role of early probiotic administration in promoting gut health and immune development. While 16S rRNA gene sequencing offers taxonomic insights, follow-up metagenomic studies are needed to uncover the functional capabilities of early microbial communities. Methods Ethics statement All procedures outlined in this study were approved by the Institutional Animal Care and Use Committee (AUP2022-0411) at Clemson University. Experimental design Fertilized broiler eggs (Ross 308 variety) were generously provided by Fieldale Farms, GA, and transported to Morgan Poultry Center (Clemson University, SC). The eggs were incubated in a sanitized egg incubator (GQF Manufacturing Company Inc., GA) with the temperature set to 37°C and humidity maintained between 55-73%, featuring automated turning. On embryonic day 18 (ED18), the eggs were candled to remove the infertile ones. 110 eggs were randomly selected to receive in ovo probiotic administration, and the other 110 were untreated. Sexing of embryos and hatchlings were not performed, and birds used in the study were of mixed sex. Preparation of the lactobacilli culture and in ovo inoculation Poultry-specific Lactobacillus strains ( L. reuteri -P43, L. acidophilus -P42, L. animalis -P38, and L. crispatus -C25) were provided by H. M. Hassan’s Laboratory at NC State University. The following preparation of the lactobacilli cocktail was done as per Sharma et al., 2024. A loopful of each frozen Lactobacillus species was inoculated into a bacterial culture tube containing 10 mL of MRS (DeMan, Rogosa, and Sharpe) at room temperature. Tubes were incubated overnight at 37°C under anaerobic conditions in a gas jar containing anaerobic packs (BD GasPak™). After 16 hours of incubation, 500 μL of the overnight culture (1%) was inoculated into 50 mL fresh MRS broth and incubated at 37°C under anaerobic conditions for 16-18 hours. The individual tubes were then centrifuged at 4000 rpm for 10 minutes at 4 ° C, and the pellet was washed two times with 1X PBS. After centrifugation, the individual pellets were then resuspended in 1X PBS, and the optical density (OD) of each culture was measured at 600 nm (OD600) using a spectrophotometer (VWR, PA). To determine the number of colony-forming units (CFUs) of each Lactobacillus strain, individual growth formulas obtained from the previous study (Sharma et al., 2024) were utilized. All lactobacilli strains were adjusted to a concentration of 10 7 CFUs/mL in PBS, and a cocktail containing a final concentration of 10 7 CFUs/mL was used. A final volume of 100 μL was injected. Post-hatch procedure As illustrated in Figure 1 , 200 hatched chicks were divided into four groups (n=50): the chicks that received probiotics in ovo were randomly divided into two groups: Group B ( in ovo probiotic supplementation only) and Group C ( in ovo and oral probiotic supplementation), and the eggs that did not receive probiotics in ovo were randomly divided into two groups: Group A (control) and Group D (oral probiotic supplementation only). After hatching, 200 hatched chicks were transported to the Godley-Snell Research Facility (Clemson University, SC) for housing. All birds that received probiotics in ovo and post-hatch were housed in separate pens in the same room, while the birds that did not receive any treatments (Group A; control group) were housed in a separate room. Each group of birds was divided into three replicates to minimize the cage effect (n=16-17 per replicate). Chick rearing was done according to the Ross 308 broiler management guidelines by Aviagen®. The birds were fed a standard diet (Purina® Start & Grow® Non-Medicated Chick Feed), and ad libitum water was provided. The birds were raised in floor pens covered with wood shavings. The birds in Groups C and D received oral gavage of the probiotic cocktail containing L. reuteri, L. crispatus, L. animalis , and L. acidophilus (10 6 CFUs/mL) on the first day of hatch, and then weekly up to five weeks of age. Sample collection Weekly euthanasia procedures involved randomly selecting and euthanizing ten chickens from each group using CO2 euthanasia over a time period of five weeks. Cecal contents were collected from the ceca in 2 mL tubes and kept in a -80°C freezer (8 samples/group/week). Six cecal tonsils were collected from each group weekly in RNA later and stored in a -20°C freezer for RNA extraction. DNA extraction Total cecal DNA was extracted from 300-400 μL cecal content using the NucleoSpin® DNA soil kit (Macherey-Nagel, Germany). The DNA mass and purity were measured using the NanoDrop One Spectrophotometer (Thermofisher Scientific, MA), ensuring that the 280/260 and 260/230 ratios of the genomic DNA fell within the range of 1.8-2. RNA isolation and cDNA preparation Cecal tonsils were homogenized using the Bead Ruptor Elite (Omni International, GA) for RNA extraction. TRIzol TM (Invitrogen, USA) was employed as per the manufacturer’s instructions to extract total RNA, followed by DNase treatment (DNA-free TM kit, Invitrogen, USA) to get rid of genomic DNA. The Nanodrop One spectrophotometer (Thermo Scientific, USA) was used to evaluate RNA mass and purity. Subsequently, cDNA synthesis was conducted with the Superscript® II First-Strand Synthesis Kit (Invitrogen, USA) and oligo-dT primers (Thermofisher Scientific, USA), following the manufacturer’s protocol. The resulting cDNA was diluted in nuclease-free water (Thermo Scientific, USA) at a 1:10 ratio. Quantitative real-time polymerase chain reaction (RT-PCR) RT-qPCR was performed using the LightCycler® 480 system (Roche Diagnostics). The PCR master mix included 10 µL of SYBR TM Green Master Mix (PowerTrackTM, USA), 1 µL each of forward and reverse primers (10 µM), and 3 µL of nuclease-free water. Each reaction comprised 15 µL of the master mix and 5 µL of cDNA in a 96-well PCR plate (USA Scientific, USA). The PCR cycling protocol involved an initial denaturation step at 95°C for 5 minutes, followed by 45 amplification cycles. Each cycle consisted of 10 seconds at 95°C for denaturation, annealing (according to the target primers specified in Table 1 ), and extension at 72°C for 10 seconds. All primers used in this study were synthesized by Sigma-Aldrich (St. Louis, MO). The mRNA expression levels of the target genes were normalized to the housekeeping gene (β-actin) using the Roche LightCycler 480 software, based on the 2 -ΔΔCT method (Livak & Schmittgen, 2001). View this table: View inline View popup Table 1 Primer sequences used for real-time quantitative PCR. 16S rRNA library preparation Genomic DNA was diluted to a concentration of 10 ng/µL per well in a volume of 10 µL in a 96-well PCR plate and submitted to the Genomic Sciences Laboratory at North Carolina State University in Raleigh, NC, for library preparation. The V3-V4 hypervariable region of the 16S rRNA gene of the bacterial DNA was amplified using the following forward primer pairs: 16SF: 5’ -TCG TCG GCA GCG TCA GAT GTG TAT AAG AGA CAG CCT G GGN GGC WGC AG - 3’ and 16SR: 5’ - GTC TCG TGG GCT CGG AGA TGT GTA TAA GAG ACA GGA CTA CHV GGG TAT CTA ATC C - 3’. The PCR cycling conditions included an initial denaturation step at 95°C for 3 minutes, followed by 25 cycles of denaturation at 95°C for 30 seconds, annealing at 55°C for 30 seconds, and extension at 72°C for 30 seconds. A final extension step was performed at 72°C for 5 minutes. Following the amplification cycles, the reaction mixture was held at 4°C until further processing. After PCR amplification, the PCR products were purified using AMPure XP bead cleanup (Beckman Coulter Life Sciences). The DNA samples were quantified and normalized to a loading library concentration of 4 nM. The amplicons were subjected to paired-end sequencing on an Illumina MiSeq platform using the MiSeq V3 300x2 sequencing kit. The sequencing was performed with ≥ 25% PhiX control spike in. Paired-end reads of all FASTQ files are available in the Sequence Read Archive (SRA) under BioProject PRJNA1205772. Bioinformatic analysis A total of 26,212,889 raw sequencing reads were generated in fastq.gz format. Demultiplexing was performed using bcl2fastq2 Conversion Software v2.20 (Illumina). Raw sequences were processed using mothur v.1.45.2, following standard operating protocol (Kozich et al., 2013). Paired-end reads were merged into contigs, quality-filtered (maxambig = 0, maxlength = 475, maxhomop = 8), and aligned to the SILVA reference database v132 trimmed to the V3–V4 region. Sequences were then dereplicated and pre-clustered. Chimeric sequences were identified and removed using VSEARCH algorithm. Taxonomic classification was performed using the RDP classifier with the RDP training set v16. Lineages associated with mitochondria, chloroplasts, unknown, Archaea, and Eukaryota were removed. A total of 459,228 Amplicon Sequence Variants (ASVs) were generated, of which low-abundance ASVs (<5 reads in the total dataset) were filtered out, retaining 16,489 ASVs for downstream analysis. After preprocessing the data in mothur, phyloseq (v.1.48.0), ggplot2 (v. 3.5.1), dplyr (v.1.1.4) and vegan (v.2.6) packages in R software (v.4.4.0) were used. Alpha diversity was assessed using the Shannon index. Kruskal-Wallis test was used for initial comparison among groups, using a Dunn’s post hoc test. Beta diversity was assessed using the Bray-Curtis dissimilarity for differences in microbial community composition across groups from the vegan R package (Oksanen et al., 2024). PERMANOVA was conducted using the adonis function from the vegan package in R. Multivariable Association with Linear Models (MaAsLin2) was used to calculate differentially abundant ASVs across treatment groups within each week (Mallick et al., 2021). Heatmaps of the relative abundances of dominant ASVs at the family level were created using the pheatmap R package, using a log 10 transformation before plotting relative abundance. Gene expression analysis For the gene expression study, data were analyzed, and graphs were created using R software (v.4.4.0). A non-parametric test (Kruskal-Wallis test) was used, and the results were considered significant if P<0.05. Data are shown graphically as the mean of the relative gene expression data (2 −ΔΔCt ) ± the standard error of the mean (SEM). Data availability statement The 16S rRNA sequencing data generated during this study have been deposited in the NCBI Sequence Read Archive (SRA) under BioProject accession number PRJNA1205772. Competing interests The authors declare no competing interests. Funding Declaration This research was supported by the USDA National Institute of Food and Agriculture Hatch Project SC-1700628 (Accession Number 7004405, Technical Contribution No. 7456), South Carolina Department of Agriculture (ACRE CGP) and Clemson University’s R-Initiatives for funding. Acknowledgments The authors also acknowledge Fieldale Farms Corporation for providing fertilized Ross 308 broiler eggs and the staff at Godley Snell Research Facility and Morgan Poultry Center for assistance with bird rearing. Data analysis was conducted using Clemson University’s Palmetto HPC System, which was made possible with support from the Clemson University Genomics and Bioinformatics Facility (CUGBF), which receives support from the College of Science and two Institutional Development Awards (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant numbers P20GM146584 and P20GM139769. Funder Information Declared South carolina department of agriculture USDA National Institute of Food and Agriculture References 1. ↵ Kogut , M. H . The effect of microbiome modulation on the intestinal health of poultry . Anim Feed Sci Technol 250 , 32 – 40 ( 2019 ). OpenUrl CrossRef 2. ↵ Taha-Abdelaziz , K. , Hodgins , D. C. , Lammers , A. , Alkie , T. N. & Sharif , S . Effects of early feeding and dietary interventions on development of lymphoid organs and immune competence in neonatal chickens: A review . Vet Immunol Immunopathol 201 , 1 – 11 ( 2018 ). 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Share Early-Life Poultry-Derived Lactobacilli Drive Microbial Succession and Gut Immune Modulation in Broiler Chickens Shreeya Sharma , Anna Seekatz , Mohammadali Alizadeh , Hosni Hassan , Alexander Yitabrek , Scott Pratt , Khaled Abdelaziz bioRxiv 2025.08.01.668251; doi: https://doi.org/10.1101/2025.08.01.668251 Share This Article: Copy Citation Tools Early-Life Poultry-Derived Lactobacilli Drive Microbial Succession and Gut Immune Modulation in Broiler Chickens Shreeya Sharma , Anna Seekatz , Mohammadali Alizadeh , Hosni Hassan , Alexander Yitabrek , Scott Pratt , Khaled Abdelaziz bioRxiv 2025.08.01.668251; doi: https://doi.org/10.1101/2025.08.01.668251 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Microbiology Subject Areas All Articles Animal Behavior and Cognition (7629) Biochemistry (17660) Bioengineering (13881) Bioinformatics (41909) Biophysics (21435) Cancer Biology (18576) Cell Biology (25479) Clinical Trials (138) Developmental Biology (13366) Ecology (19887) Epidemiology (2067) Evolutionary Biology (24301) Genetics (15598) Genomics (22482) Immunology (17726) Microbiology (40359) Molecular Biology (17162) Neuroscience (88529) Paleontology (666) Pathology (2830) Pharmacology and Toxicology (4820) Physiology (7636) Plant Biology (15125) Scientific Communication and Education (2044) Synthetic Biology (4290) Systems Biology (9817) Zoology (2269)