Recruitment and Establishment of the Gut Microbiome in Arctic Shorebirds Kirsten Grond1,∗, Richard B Lanctot2, Ari Jumpponen1 and Brett K Sandercock1

FEMS Microbiology Ecology, 93, 2017, fix142

doi: 10.1093/femsec/fix142 Advance Access Publication Date: 24 October 2017 Research Article

RESEARCH ARTICLE Recruitment and establishment of the gut microbiome in arctic shorebirds Kirsten Grond1,∗, Richard B Lanctot2, Ari Jumpponen1 and Brett K Sandercock1

1Kansas State University, Division of Biology, Manhattan, KS 66506, USA and 2US Fish and Wildlife Service, Migratory Bird Management, Anchorage, AK 99503, USA

∗Corresponding author: Department of Molecular and Cell Biology 91 North Eagleville Road University of Connecticut Storrs, CT 06269-3125, USA. Tel: +7854776545; E-mail: [email protected] One sentence summary: Shorebird embryos hatch with negligible gut microbiota, and chicks acquire a stable bacterial community within three days of hatching Editor: Cindy Nakatsu

ABSTRACT

Gut microbiota play a key role in host health. Mammals acquire gut microbiota during birth, but timing of gut microbial recruitment in birds is unknown. We evaluated whether precocial chicks from three species of arctic-breeding shorebirds acquire gut microbiota before or after hatching, and then documented the rate and compositional dynamics of accumulation of gut microbiota. Contrary to earlier reports of microbial recruitment before hatching in chickens, quantitative PCR and Illumina sequence data indicated negligible microbiota in the guts of shorebird embryos before hatching. Analyses of chick feces indicated an exponential increase in bacterial abundance of guts 0–2 days post-hatch, followed by stabilization. Gut communities were characterized by stochastic recruitment and convergence towards a community dominated by Clostridia and Gammaproteobacteria. We conclude that guts of shorebird chicks are likely void of microbiota prior to hatch, but that stable gut microbiome establishes as early as 3 days of age, probably from environmental inocula.

Keywords: 16S rRNA gene; bacteria; Calidris; gut microbiota; qPCR; precocial young

INTRODUCTION initial composition of gut microbiota in mammalian offspring (Stevens and Hume 1998). Gut microbiota contribute to maintaining organismal health It remains unclear whether birds can acquire gut microbiota through nutrient uptake (Leser and Mølbak 2009), detoxification while inside the egg, and how microbial communities accumu- of digestive byproducts (Kohl 2012), energy and fat metabolism late after initial recruitment. Bacterial colonization can theo- (Velagapudi et al. 2010), and interactions with the host immune retically occur before or after hatching, and we propose two system (Rescigno 2014). Environmental exposure early in life alternative hypotheses to describe the process of microbial col- can shape the gut microbiota and aid in defending against onization: the ‘head start’ and the ‘sterile egg’ hypothesis. The pathogens while the immune system is immature (Bar-Shira, ‘head start’ hypothesis posits that microbes enter the gastroin- Sklan and Friedman 2003). In mammals, maternal vaginal and testinal tract of avian embryos through transovarian transmis- fecal microbe transmission are crucial in the recruitment and sion during oogenesis as a maternal effect, or possibly from establishment of microbiota in the digestive tract of a neonate the environment by penetration through eggshell pores and (Palmer et al. 2007). Thus, maternal effects strongly control the

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embryonic membranes after egg laying (Gantois et al. 2009;Cox ples collected from shorebird chicks from hatch to up to 10 days et al. 2012; Martelli and Davies 2012). The ‘sterile egg hypoth- of age. esis’ predicts that microbiota recruitment in the avian gut occurs after hatch because the chorion membrane maintains a sterile environment within the egg (van der Wielen et al. 2002; MATERIALS AND METHODS Kohl 2012). The development of the avian embryo within the Sample collection and preparation membrane-enclosed yolk supports the latter hypothesis, but enteric bacteria have been cultured from the guts of embryos of do- We collected 27 eggs from seven clutches of dunlin (Calidris mestic chicken (Gallus gallus domesticus; Kizerwetter-Swida´ and alpina) and 18 eggs from five clutches of semipalmated sand- Binek 2008), and vertical transfer of pathogenic Salmonella enter- pipers (Calidris pusilla), at Utqiagvik˙ (formerly Barrow), Alaska ◦ ◦ ica has been documented between mother and embryo in do- (71 17 26 N, 156 47 19 W) in June–July 2013. We estimated incu- mestic chickens (Guard-Petter 2001; De Buck et al. 2004). The two bation stage via changes in egg buoyancy that occur with em- hypotheses differ in the maternal contribution to gut microbiota bryonic development (Liebezeit, Smith and Lanctot 2007)and of offspring and in the source of inoculum. The ‘head start’ hy- collected eggs 1–3 days before the predicted hatch date to en- pothesis predicts that maternal or environmental inoculation sure near complete embryo development. After collection, em- occurs before hatching, possibly resulting in metabolic and im- bryos were removed from eggs, euthanized and kept frozen at ◦ munological advantages at hatching. In contrast, if gut micro- –20 C until dissection. Before dissection, embryos were washed biota are absent in embryos, as posited by the ‘sterile egg’ hy- in a weak solution of 0.6% sodium hypochlorite (10% bleach) to pothesis, chicks must recruit microbiota from the environment minimize contamination by external microorganisms. We asep- after hatch. tically removed the lower intestinal tract between the gizzard ◦ Developmental strategies after hatching vary within birds and cloaca and stored samples at –80 C. We also collected the and range from chicks hatching with complete dependence invaginated yolk sacs from the embryos of five dunlin and four on their parent(s), like most altricial species, to superprecocial semipalmated sandpipers. species that are independent of parental care after hatching. Al- To investigate gut microbiota of mobile broods after hatch- tricial and precocial birds likely differ in the degree of mater- ing, we selected dunlin and red phalaropes (Phalaropus fuli- nal influence on microbial inoculation of their chicks. Anal- carius) as study species because dunlin use mesic terres- tricial bird species, the Barn Swallow (Hirundo rustica)showed trial habitat, whereas red phalaropes use freshwater habitat slight maternal, but not paternal, effects on the fecal micro- (Cunningham, Kesler and Lanctot 2016). Different habitats and biota of their offspring (Kreisinger et al. 2017). Altricial birds have associated microbiomes could affect the recruitment of gut mi- undeveloped young that require parental feeding and brooding crobiota in chicks. We fit the attending adults with 1.55 g very until fledging, whereas precocial chicks hatch fully developed high frequency (VHF) radios, and, for dunlin, one chick per brood with invaginated yolk sacs, leave the nest immediately, forage with 0.26 g VHF radios (Holohil Systems Ltd., Carp, Ontario, independently, but rely on parental brooding and defense un- Canada; Fig. S3.1, Supporting Information). Red phalarope chicks til able to thermoregulate (Sibly et al. 2012). Thus, altricial birds were too small at hatch to apply transmitters so we located have higher potential for parental influence on gut microbiota young by tracking the attending male. We attempted to locate through salivary transfer and a longer period of nest occupa- mobile broods every 3 days, using radio telemetry and by search- tion, whereas precocial parents have little direct influence on ing areas where broods were last seen. Once broods were lo- their offspring’s microbiota aside from exposure to bacteria on cated, chicks were captured and feces were collected by placing feathers during brooding, and leading young to brood-rearing the chicks in individual compartments in an insulated, ther- areas. mally heated bag, lined with sterile wax paper. Chicks were held In addition to uncertainty about the timing of initial recruit- for <5 min to minimize stress and then released together. At ment of microbiota, the early successional dynamics of the avian several capture sites, we also collected environmental samples gut microbiota are unclear. For example, it remains unknown consisting of a mix of water and soil in a 1.5-ml Eppendorf tube how rapidly chick guts are colonized, in what order bacterial using a flame-sterilized spatula. All samples were stored in 100% ◦ taxa become established and when convergence towards an ethanol at –20 C until further analyses. adult gut microbiome occurs. Precocial young of most Arctic- To remove ethanol from fecal samples, we centrifuged sam- breeding shorebirds are nidifugous, self-feeding and dependent ples for 10 min at 10 000 rpm and discarded the supernatant. We on parents for thermoregulation, and could benefit from recruit- repeated this step twice with 1 ml of RNase/DNA-free molec- ing gut microbiota before hatching, because symbionts can pro- ular grade water to minimize ethanol contamination (Grond mote nutritional uptake and growth immediately after hatch et al. 2014;Ryuet al. 2014). We shredded embryonic gastroin- (Angelakis and Raoult 2010). Efficient use of nutrients may be testinal tissues, and extracted DNA from embryo and chick fecal particularly important in the Arctic because a short and syn- samples using the MoBio Power Lyzer/Power Soil kit following chronous breeding season requires rapid chick development the manufacturer’s instructions (Mo Bio Laboratory, Carlsbad, (Schekkerman et al. 2003), favoring the recruitment of microbiota CA, USA), except for replacing the bead beating step with 15 via the ‘head start’ hypothesis. min vortexing at high velocity for tissue homogenization. Yield We investigated the ontogeny of the gut microbial commu- of genomic DNA was determined using a spectrophotometer nity in three species of Arctic-breeding shorebirds. Our objec- (NanoDrop 2000, Thermo Fisher Scientific, Waltham, MA, USA). tives were to (i) test the ‘head start’ and ‘sterile egg’ hypotheses and determine if precocial chicks of arctic shorebirds recruit Conventional PCR gut microbiota before or after hatching, and (ii) assess successional trajectories in the gut microbiota of shorebird chicks af- To test for the presence of bacteria in the embryonic gut, we ter hatching. To address our two objectives, we combined quan- PCR-amplified total bacterial communities using general bacte- titative PCR (qPCR) and Illumina MiSeq techniques to analyze rial primers 515F and 806R to generate 16S rRNA gene ampli- the gut microbiota of embryos prior to hatch and in fecal sam- cons (Caporaso et al. 2012). Primer sequences are provided in the Grond et al. 3

Supplemental Materials (Table S3.1, Supporting Information). et al. 2006). For full QIIME pipeline, we refer to the QIIME Code PCR reactions were conducted in a 25-μl reaction volume, using supplement. We identified chimeras—artifacts that combine AmpliTaq Gold DNA polymerase (Applied Biosystems, Waltham, multiple different sequences—using CHIMERASLAYER (Haas MA, USA) and 5 μl of DNA template (5 ng-DNA/μl). PCR condi- et al. 2011). Sequences were clustered to operational taxonomic tions consisted of 30 cycles of 15 s at 95◦C, 30 s at 55◦Cand30s units (OTUs) at 97% sequence similarity, and assigned to taxa us- at 72◦C, preceded by an initial denaturing step of 10 min at 95◦C, ing the Naive Bayesian Classifier with an RDP reference (Wang and followed by a final extension step of 5 min ◦at 72 C. et al. 2007). After taxonomy assignment, we identified and To ensure that the bacterial DNA within the embryo lower removed non-target sequences of chimeras, singletons, non- gastro-intestinal (GI) tract was not degraded by our disinfection aligned sequences, archaea, chloroplasts and mitochondria. procedures, we amplified avian DNA from 24 random samples Prior to subsequent analyses, we rarefied chick samples to with primers 2550F and 2718R designed for molecular sexing of 5000 sequences per sample due to low-sequence yields in sam- non-ratite birds, following PCR conditions of Fridolfsson and El- ples of chicks from 0–2 days of age. All embryo samples yielded legren (1999). Had the sodium hypochlorite compromised the far fewer than 5000 sequences, and were analyzed without DNA, we should not have been able to acquire avian or bacte- rarefication. rial amplicons. Validation tests confirmed the integrity of the lab procedure and our ability to amplify the bacterial DNA. STATISTICAL ANALYSIS Quantitative PCR Embryo gut and yolk microbiota

We estimated 16S rRNA gene copy numbers in embryo gut, yolk, Statistical analyses were conducted in R (ver. 3.3.1, R Develop- fecal and control samples in triplicate qPCR reactions on a Bio- ment Core Team 2016). To test for differences in 16S rRNA gene Rad CFX96 Touch Real-Time PCR thermocycler (Bio-Rad Labora- copy numbers between embryos and chicks, we performed a tories, Inc., Berkeley, CA, USA). We used a TaqMan R qPCR assay one-way analysis of variance (ANOVA) and assessed pairwise (Applied Biosystems) targeting the bacterial 16S rRNA gene in differences using a post hoc Tukey HSD test with functions of combination with 2X TaqMan Gene Expression Master Mix with the base Program R. We compared bacterial richness (number of two general bacterial primers at 100 nM final concentrations OTUs) among our embryo gut, yolk and negative controls using (F˙Bact1369 and R Prok1492; Furet et al. 2009; primer sequences an ANOVA, followed by a post hoc Tukey HSD test. To compare μ can be found in Table S1). Cocktails included 4 l of DNA tem- microbial communities of embryo gut and yolk samples and the μ ◦ plate(5ng-DNA/ l) and conditions consisted of 2 min at 50 C negative control, we applied non-metric multidimensional scal- and an initial denaturing for 10 min at 95◦Cfollowedby40cy- ◦ ◦ ing (NMDS) of Bray-Curtis distance matrices using the metaMDS cles of 15 s at 95 C and 1 min at 60 C. The negative controls were function in the ‘vegan’ package. RNase/DNA-free molecular grade water and the positive controls was Semipalmated Sandpiper fecal DNA at 5 ng-DNA/μl. We included our positive and negative controls in triplicate in Chick gut microbiota the two qPCR runs we conducted. Standard curves were generated using 2 to 2 × 106 16S rRNA gene copies of Staphylococcus To look for natural breaks in bacterial abundance in our chick aureus subsp. aureus (efficiency = 105–116%, R2 = 0.984–0.990). To qPCR data, we used the segmented function from the ‘Segmented’ control for the possibility of PCR inhibition and to establish min- package (Muggeo 2008). Large changes in the slope represent imum detection levels, we spiked a subset of samples (n = 15) changes in the accumulation of 16S rRNA copies in the chick gut with a dilution series of the S. aureus rRNA gene ranging from and were used to determine whether we could pool our chick 100 to 106 copies. microbiota data into naturally occurring broader age classes. To test for significant differences among slopes in bacterial abun- Illumina sequencing and sequence analyses dance versus chick age, we ran a Davies test using the davies.test function from the ‘Segmented’ package. We used general bacterial 515F and uniquely barcoded 806R We calculated diversity indices (observed number of primers to generate multiplexed 16S rRNA gene amplicons from OTUs, Simpson’s (1-D) and Shannon’s H) using the QIIME embryo guts and fecal samples, following protocols of the Earth alpha diversity.py script. In addition, we calculated evenness Microbiome Project (Caporaso et al. 2012). PCR reactions were (Pielou’s J) using the diversityresult function in the ‘BiodiversityR’ performed in a 25-μl reaction volume, using AmpliTaq Gold DNA package (Kindt and Coe 2005). We estimated beta diversity to polymerase (Applied Biosystems) and 5 μl of DNA template (5 compare variance between age groups (0–2 days versus 3–10 ng-DNA/μl), and were run using 25 cycles, opposed to the 35 cy- days) and environment. We tested for pairwise differences cles described in the Earth Microbiome Project (EMP) protocols. among age groups using post hoc Tukey HSD tests. We removed primers from our PCR product using the Agencourt We determined shared core OTUs among young and old AMPure XP PCR purification system (Beckman Coulter, Brea, CA) chicks and environmental samples, and constructed a scaled following the manufacturer’s instructions except AMPure vol- Venn diagram using EulerAPE (Micallef and Rodgers 2014). Core ume ratio was adjusted to 1:1 and the ethanol washes were re- OTUs were defined as OTUs that were present in >50% of our peated three times instead of two. We sequenced the V4 region samples within a group, and were determined using the QIIME in 2 × 250 bp paired-end runs using the Illumina MiSeq platform. compute core microbiome.py script. We used the core commu- Each Illumina run included a 15% PhiX spike. Embryo samples nity instead of the entire bacterial community as peripheral low and fecal samples from chicks of 0–3 days old were sequenced abundance OTUs could skew our results. twice because of low-sequence yields. To investigate potential differences among microbial com- Bacterial 16S rRNA gene sequences were quality filtered, con- munities in chicks and the environment, we applied NMDS. tiged and demultiplexed using the QIIME program (Caporaso We tested for treatment differences through an analysis of et al. 2010), and aligned against the GreenGenes 16S rRNA gene similarity using the anosim function in the ‘vegan’ package. reference database using PyNAST algorithms (v.13 8;DeSantis Means are shown ± standard error. 4 FEMS Microbiology Ecology, 2017, Vol. 93, No. 12

To determine whether variation in gut communities differed among samples from dunlin and red phalarope chicks, or if samples could be pooled for further analyses, we analyzed homo- geneity of variance using the betadisper function from the ‘vegan’ package (Oksanen et al. 2015). To test for pairwise differences in variance between young and old chicks and environmental samples, we performed a permutation F test using the permutes function in the ‘vegan’ package. Because we sampled multiple chicks within the same brood, we tested for potential effects of brood identity on OTU richness and bacterial abundance (log10 16S rRNA copy numbers). We compared Akaike Information Criterion (AIC) values generated by linear mixed effect models using the lmer and glm function in the ‘vegan’ and ‘lme4’ packages (Bates et al. 2015;Oksa- Figure 1. Median (thick, black line), 25% and 75% quartiles (boxes), and 90% con- nen et al. 2015). Our null model included a fixed effect of chick fidence intervals (whiskers) for the log10 16S rRNA gene copy numbers. (a)Sam- ples included lower gastrointestinal tracts (gut) and yolk sacs from developed age, and our full model included an additional random effect of embryos of dunlin (DUNL) and semipalmated sandpipers (SESA) at Utqiagvik,˙ Brood ID. Alaska, 2013. The negative control was RNase/DNA-free molecular grade water, whereas the positive control was a fecal sample from an adult SESA (adult (+); Data accessibility 5ngDNA/μl). Dashed lines and letters above boxes represent significant differ- The nucleotide sequences and metadata have been made avail- ences in 16S rRNA copy number (Tukey HSD). The microbial community from able through Figshare at https://doi.org/10.6084/m9.figshare. the positive control was not included in further analyses, but is shown here for 4587394.v1 and https://doi.org/10.6084/m9.figshare.4587379.v1. reference purposes. (b) 16S rRNA gene copy numbers in fecal samples from REPH and DUNL chicks from 0–10 days after hatch. RESULTS significantly different from the negative control (F = 101.6, Timing of recruitment 2,38 P = 0.13), and richness of embryo gut samples did not differ Conventional and qPCR from yolk samples (TukeyHSD, P = 0.71). Our NMDS showed Conventional PCR assays yielded no visible 16S rRNA gene am- that the minimal number of sequences obtained samples from plification from gut or yolk samples of shorebird embryos col- Semipalmated Sandpiper and Dunlin embryos, and our negative lected 1–3 days before hatching. Consistently, the high fidelity control overlapped in ordination space (Fig. S2, Supporting In- and high sensitivity Taqman qPCR assays suggested low copy formation). Overall, our sequence data indicated that minimal numbers of 16S rRNA genes in the intestinal tract and yolk sac of microbiota were present in the gastrointestinal systems or yolk embryos. The qPCR detected bacterial amplification in our pos- sacs of shorebird embryos before hatching of the eggs. itive control after 12.6 ± 1.1SE cycles, compared to 33.8 ± 0.2 and 33.2 ± 0.3 cycles for the guts of dunlin and semipalmated Establishment of the microbiota sandpiper embryos, 34.1 ± 0.2 and 33.7 ± 0.2 cycles for yolk samples, and 33.9 ± 0.5 cycles for our negative control. We detected We collected 45 fecal samples from 1–10 day old chicks of dun- negligible copy numbers of the bacterial 16S rRNA gene in all lin and red phalaropes at Utqiagvik,˙ AK, in 2013. Despite the dif- embryo samples, and we pooled samples from semipalmated ferent habitats used by dunlin and red phalaropes to rear their sandpipers and dunlin for statistical analyses. Our gut and yolk chicks, fecal samples of chicks did not differ in the abundance = = samples did not differ from the negative controls in either and variance of their microbiota (F2,27 5.5, P 0.16). We pooled threshold cycle or copy numbers (Tukey HSD; P > 0.22; Fig. 1a), samples for further analyses to increase sample sizes. We did and all samples and negative controls had higher threshold cy- not detect an effect of brood identity on the bacterial abun- = = cle and lower copy numbers than our positive controls (Tukey dance and OTU richness (AICnull 285.6 and 66.7; AICfull 283.6 HSD; P < 0.001). and 68.7), and we therefore regarded samples of chicks from the Avian DNA from all 24 gut samples successfully amplified in same brood as independent. conventional PCR, further confirming that disinfecting the embryo’s external surfaces did not compromise DNA within the GI Abundance tract of shorebird embryos. We detected 10 copies of the 16S We observed an increase in 16S rRNA gene copy numbers from rRNA of S. aureus after 33.7 ± 0.21 cycles, which was a similar age 0–2 days. After 3 days post hatch, copy numbers stabilized detection threshold as found for our embryo and negative con- at on average 395 852 ± 98 048 per sample (Fig. 1b). Our broken- trol samples. Thus, our conventional and qPCR assays indicate stick regression indicated a natural break at 3 days post hatch, that low copy numbers of bacterial rRNA were present in our and the slopes between 0–2 and 3–10 days post hatch differed embryo samples, which represented the entire gastrointestinal significantly (Davies’ test, P < 0.001; Fig. S3, Supporting Informa- system from the gizzard to cloaca. tion). The slope from day 3–10 did not differ from 0, suggesting stability with no change in microbial abundance over this period Sequencing (t = 0.095, P = 0.925). As a result, we pooled our samples into a Illumina MiSeq libraries acquired from 42 of our 45 embryo ‘young’ (0–2 day age) and ‘old’ (3–10 day age) group for further samples contained an average of 108.6 ± 15.9 sequences (range analyses. 0–427), whereas the positive control contained over 7 million sequences. Three samples did not provide any sequencing out- Richness and composition put and were not included in further analyses. A second se- After quality filtering, we retained 1689 081 high-quality sequence library generated from the gastrointestinal tracts and quences (90.2%) from our total of 1860 751 raw sequences yolk samples from shorebird embryos resulted in similarly low for further analyses. After rarefaction to 5000 sequences per yields. The number of OTUs in the embryo samples was not sample, we included five samples of 0- to 2-day-old chicks, Grond et al. 5

Figure 3. NMDS of the OTUs in the fecal samples of young chicks (0–2 days, n = 5) and old chicks (3–10 days, n = 19) of dunlin and red phalaropes, and environmental samples (n = 6), at Utqiagvik,˙ Alaska, 2013. Figure 2. Scaled Venn diagram of the core bacterial OTUs isolated from fecal samples of young (0–2 days) and old (3–10 days) dunlin and red phalarope chicks, as well as environmental samples of water and soil. Numbers represent numbers of unique or shared core OTUs.

19 samples of 3- to 10-day-old chicks and six environmental samples. Overall, environmental samples were highest in bacterial richness on all phylogenetic levels, followed by young chicks and old chicks (Table 1;Fig.2; Fig. S4, Supporting Information). We identified a total of 36, 723 and 5715 unique bacterial phyla, genera and OTUs in our samples. Observed OTU richness was lower in samples from old chicks than in samples from young chicks (Tukey HSD, P = 0.03), consistent with environmental/habitat filtering occurring as chicks aged. In both young and old chicks, the observed OTU richness was lower than in the environmental samples (Tukey HSD, P < 0.001). Simpson’s (1-D) and Shannon’s (H) diversity indices indicated that environmental samples had Figure 4. Relative abundance of bacterial phyla in environmental samples and the highest diversity scores (0.98 ± 0.01; 8.10 ± 0.39), and old fecal samples of young (0–2 days) and old (3–10 days) chicks of dunlin and red chicks the lowest (0.56 ± 0.04; 1.98 ± 0.14). In addition, evenness phalaropes at Utqiagvik,˙ Alaska, 2013. Day 0 represents day of hatching. of bacterial communities in the gut was highest in environmental samples (0.81 ± 0.03), followed by young chicks (0.53 ± 0.11) dance decreased from 45.5 ± 14.4% in young to 22.3 ± 6.0% in and old chicks (0.30 ± 0.02; Fig. S4). old chicks. Within Proteobacteria, 43% of sequences belonged to At a community level, our NMDS (k = 3, stress = 0.083) distin- the genus Rickettsiella within the Gammaproteobacteria.Therel- guished environmental from chick samples (ANOSIM, R = 0.853, ative abundance of Actinobacteria and Bacteroidetes declined by P = 0.003; Fig. 3). Young chick sample communities differed 50%–80% from young to old chicks (Actinobacteria:4.1± 1.5% to from old chick communities (ANOSIM, R = 0.487, P = 0.006), 0.3 ± 0.1%; Bacteroidetes:5.0± 2.2% to 0.1 ± 0.02%; Fig. 5). but were not significantly different from environmental samples Beta diversity of bacterial OTUs was greater among young (ANOSIM, R = –0.052, P = 0.599). However, microbial communi- chicks than among the environmental samples (F2,27 = 3.347, ties of old chick samples were highly differentiated from envi- P = 0.03), but did not differ between old chicks and the environmental samples (ANOSIM, R = 0.999, P < 0.001). ronment (F2,27 = 3.347, P = 0.26), and only marginally between The gut microbiota composition of chicks was dynamic young and old chicks (F2,27 = 3.347, P = 0.07). over time. Relative abundance of Firmicutes increased from 37.7 ± 14.2% in young to 73.4 ± 5.9% in old chicks (Fig. 4). The increase was mainly driven by increases in abundance of the DISCUSSION classes Clostridia and Bacilli (Fig. 5). Within the class Clostridia, Timing of recruitment Clostridium colinum (sequence similarity = 98%; Genbank Ac- cession LC011040.1)—an opportunistic avian pathogen—was re- Our data largely supported the ‘sterile egg hypothesis’, which sponsible for 73.6 ± 8.6% of sequences among young chicks and posits that embryos hatch from eggs with a sterile gut. De- 88.1 ± 4.1% in old chicks. The second most abundant OTU within spite detecting low copy number of 16S rRNA genes in embryo the Clostridia belonged to the genus Candidatus Arthromitus with samples using conventional and qPCR assays, these were either a relative abundance of 6.1 ± 2.7% in young and 10.3 ± 4.1% in below detection level (conventional PCR) or indistinguishable old chicks. In contrast to Firmicutes, Proteobacteria relative abun- from negative controls to which no DNA template had been 6 FEMS Microbiology Ecology, 2017, Vol. 93, No. 12

Table 1. Total and per sample bacterial richness at different phylogenetic levels in guts of young chicks (young; 0–2 days old), old chicks (old; 3–10 days), and in environmental samples (environment).

Total phyla Phyla/sample Total genera Genera/sample Total OTU OTU/sample

Young 23 16.2 ± 1.5 432 132.2 ± 32.9 1279 308.4 ± 80.1 Old 19 9.5 ± 0.4 379 42.5 ± 3.4 1005 105.7 ± 7.7 Environment 34 25.2 ± 1.2 568 304.7 ± 23.1 4116 1060.5 ± 122.7

Figure 5. Sequence abundance per bacterial class for the four main phyla found in environmental samples of water and soil, and young (0–2 days) and old (3–10 days) chick fecal samples collected from dunlin and red phalaropes at Utqiagvik,˙ Alaska, 2013.

added. The low copy numbers detected in the negative controls barcode switching in Illumina sequencing studies can present likely represent artifactual sequences, as products appeared late a substantial source of error. Sinclair et al. (2015) used both for- in the amplification process after >30 cycles (Arikawa et al. 2008). ward and reverse barcoded primers and estimated that barcode Similar to the PCR assays, we documented little evidence of switching could represent up to 22% of sequences in the Illumina bacterial presence in our embryo samples or the negative con- platform. We used only reverse-barcoded primers and were un- trols in the Illumina MiSeq sequencing analyses. The detection able to estimate or detect barcode switching. Sequences in neg- of any sequences in our negative controls might be attributed ative controls (or samples without adequate template) may also to several potential causes. Carlsen et al. (2012) demonstrated originate earlier in the library preparation process. Last, DNA ex- tag switching in 0.7%–1.6% of sequences during pyrosequencing, traction kits and PCR reagents may contain some bacterial DNA, resulting in erroneous assignment of sequences to samples. In leading to false detections in low template samples like our em- fact, the total number of sequences in our embryos and nega- bryos (Salter et al. 2014). tive controls comprised only 0.06% of all sequences, a proportion The low abundance of bacteria in the guts of shorebird far below error rates estimated by Carlsen et al. (2012). Similarly, embryos collected under natural conditions contrasts with Grond et al. 7

past reports from domestic chickens. Kizerwetter-Swida´ and stabilization at 3–10 days of age. Community filtering for spe- Binek (2008) cultured a variety of enteric bacteria from chicken cific, high abundance gut symbionts was supported byade- embryos obtained from a local hatcher. However, if the gut mi- crease in evenness, which suggested a rapid increase in abun- crobiota recruitment and establishment were similar in shore- dance of dominant bacterial taxa with age. Depletion of the yolk birds and chickens, we should have detected significantly higher sac after 2–3 days and the associated decrease in available nutri- bacterial loads with our culture-independent methods that tar- ents to the chick gut microbiota might also explain the decrease geted the entire spectrum of gut bacteria. Differences in environ- in diversity. After nutrients are depleted, only bacteria that use mental conditions between domestic chickens in a commercial- nutrients derived from the chick diet, or those produced by scale production facility and Arctic-breeding birds under field other gastrointestinal bacteria, would be sustained. After 3 days conditions might lead to different routes for bacterial transmis- post hatch, we observed establishment of a stable abundance sion. For example, Salmonella enteritidis can infect chicken em- of gut microbiota. Bacterial abundance did not change over the bryos via infection of the female chicken’s reproductive organs, rest of our sampling period, but then differed by two orders of resulting in incorporation of bacteria into the egg during ooge- magnitude between 10-day-old chicks and adult birds. We were nesis (Gantois et al. 2009). The likelihood of a similar infection unable to sample young birds after 10 days of age because in wild birds may be low and could possibly be the reason for chick mortality and enhanced mobility made locating chicks the limited evidence of bacteria we documented in our wild bird challenging. embryos. As chicks aged from 0 to 10 days old, their gut microbiota Another potential infection route for avian embryos is became dominated by bacteria within the phyla Firmicutes and through the egg shell and membranes, which is referred to as Proteobacteria. Increasing abundance of Clostridia was the main trans-shell infection. We detected minimal gut microbiota in driver of the rise in Firmicutes abundance. Clostridia are also early shorebird embryos gut and yolk samples, which also indicated colonizers of the human infant gastrointestinal tract and essen- that bacteria do not infect the embryo gastrointestinal tract tial in maintaining gut homeostasis (Lopetuso et al. 2013). The as a result of trans-shell infection during incubation. Whether prevalence and natural occurrence of Clostridia in the gastroin- infection of the egg after laying reduces hatching success in testinal tract in other wild birds is unknown, but their high abun- birds is still under debate (Cook et al. 2003; Wang, Firestone dance in the guts of shorebird chicks suggests a role for com- and Beissinger 2011), but incubation was shown to reduce and mensalism or potentially mutualism. change microbial communities on egg shells (Lee et al. 2014). Within the class Clostridia, C. colinum was the most abun- Since we collected our embryos close to hatch and all embryos dant species with a relative abundance of 70%. Clostridium col- were alive when the eggs were opened, trans-shell infection inum is an opportunistic avian pathogen and can cause ulcera- likely had either not occurred in our embryos or infection did tive enteritis in chickens and quail. Clostridium colinum infects not lead to reduced viability. Overall, our field data indicate that a variety of birds, including domestic poultry, northern bob- gut microbiota become established after chicks hatch and are whites (Colinus virginianus), American robins (Turdus migratorius), exposed to the environmental microbiome. western bluebirds (Sialia mexicana) and lories (Trichoglossus sp. and Eos sp.; Winterfield and Berkhoff 1977; Porter 1998; Bild- Establishment of the microbiota fell, Eltzroth and Songe 2001;Pizarroet al. 2005; Beltran-Alcrudo et al. 2008). All previous detections of C. colinum have been asso- Based on our qPCR data, the bacterial abundance increased ex- ciated with disease or death of the host, but the occurrence of ponentially in chick guts during the first 3 days after hatch- C. colinum in shorebird chicks could present evidence of alterna- ing. Precocial chicks of Arctic shorebirds leave the nest within a tive functions in the avian gut. Moreover, our sequences showed day of hatching and feed independently, and we have observed 98% similarity to C. colinum, which could indicate that our se- chicks feeding as early as 1 day of age (personal communica- quences belong to an alternative, unidentified strain of C. colinum tion: D.E. Gerik). Since parents do not provision precocial chicks or closely related Clostridium species. Clostridium colinum was not in Arctic-breeding shorebirds, chicks likely acquire gut micro- detected in the environmental samples we sequenced, suggest- biota from ingestion of prey-associated microbiota. An inocula- ing that chicks may acquire this bacterium from their diet or tion route via diet was supported by our ordination results which another source. showed that the microbiota of young chicks were more similar to Proteobacteria were the second most abundant phylum within the environmental microbiota in community composition than the chick gut; their relative abundance decreased from young old chicks. to old chicks. Similar to Clostridia, human infants experience an A rapid increase in the microbial abundance within the first early increase in the class Gammaproteobacteria (La Rosa et al. 3 days after hatching may be enhanced by the supply of yolk 2014). Gammaproteobacteria represent a variety of species, but to the chick gastrointestinal tract. During the first 2–3 days af- their functions have not been identified in avian systems. Within ter hatch, precocial chicks use an invaginated yolk sac for nutri- the Proteobacteria, we observed a shift from an evenly structured ents while they learn to forage (Forsythe 1973; Schekkerman and community including all classes of Proteobacteria to a community Boele 2009). The yolk mainly consists of fat, protein and carbo- dominated by Gammaproteobacteria. Rickettsiella was the domi- hydrates, and also contains vitamins and trace elements (Vieira nant genus within the Gammaproteobacteria. Rickettsiella spp. are 2007). Combined with the relatively high body temperature of associated with microbiota of terrestrial arthropods (Bouchon, the chick, yolk provides a rich substrate for bacterial growth, and Zimmer and Dittmer 2016; Duron, Cremaschi and McCoy 2016), is commonly used in bacterial pure culture media (Carter 1960; including several pathogens and endosymbionts (Leclerque and Westblom, Madan and Midkiff 1991;Byrneet al. 2008). Kleespies 2008; Tsuchida et al. 2010). The detection of Rickettsiella We observed higher richness at a phylum, genus and OTU in chicks could indicate an important role of diet as a microbial level among gut microbes in young chicks than old chicks. OTU inoculum. Although Rickettsiella spp. are generally classified as richness was three times higher in young chicks than old chicks arthropod pathogens (Bouchon, Zimmer and Dittmer 2016), the for some bacterial phyla, indicating either selective recruitment high relative abundance in chicks indicates previously unknown or host-controlled environmental filtering prior to microbiota non-pathogenic functions. 8 FEMS Microbiology Ecology, 2017, Vol. 93, No. 12

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