The reproductive microbiome and maternal transmission of microbiota via eggs in Sceloporus virgatus

Abstract Maternal transmission of microbes occurs across the animal kingdom and is vital for offspring development and long-term health. The mechanisms of this transfer are most well-studied in humans and other mammals but are less well-understood in egg-laying animals, especially those with no parental care. Here, we investigate the transfer of maternal microbes in the oviparous phrynosomatid lizard, Sceloporus virgatus. We compared the microbiota of three maternal tissues—oviduct, cloaca, and intestine—to three offspring sample types: egg contents and eggshells on the day of oviposition, and hatchling intestinal tissue on the day of hatching. We found that maternal identity is an important factor in hatchling microbiome composition, indicating that maternal transmission is occurring. The maternal cloacal and oviductal communities contribute to offspring microbiota in all three sample types, with minimal microbes sourced from maternal intestines. This indicates that the maternal reproductive microbiome is more important for microbial inheritance than the gut microbiome, and the tissue-level variation of the adult S. virgatus microbiota must develop as the hatchling matures. Despite differences between adult and hatchling communities, offspring microbiota were primarily members of the Enterobacteriaceae and Yersiniaceae families (Phylum Proteobacteria), consistent with this and past studies of adult S. virgatus microbiomes.


Introduction
Maternal transmission of microbiota has increasingly been documented across the animal world (Funkhouser and Bordenstein 2013, Baldo et al. 2018, Moeller et al. 2018, Youngblut et al. 2019 ), but the mechanisms of this transfer, and the heritability of the micr obiome r emains unclear and v ariable acr oss taxa (Lauder et al. 2016, Youngblut et al. 2019 ).In mammals, bacteria ar e tr ansferr ed during passa ge thr ough the birth canal, and possibly during de v elopment in the womb, although that is curr entl y under debate (Dominguez-Bello et al. 2010, Funkhouser and Bordenstein 2013, Stinson et al. 2019, Ro w e et al. 2020, Kennedy et al. 2023 ).The neonatal microbiome is then supplemented through later par ental car e, whic h can v ary widel y acr oss taxa, including nursing (Milani et al. 2017 ) and direct parent-to-offspring contact (Banning et al. 2008, Dominguez-Bello et al. 2010 ).The early inoculation of microbes is critical for healthy de v elopment of the adult microbiome, and can have long-term health consequences when perturbed (Lozupone et al. 2012, Knutie et al. 2017, Shao et al. 2019, Warne et al. 2019, Kim et al. 2020 ).
While less well-studied, there is also evidence that maternal micr obes ar e tr ansferr ed in ovipar ous animals (r e vie wed in Nyholm 2020 ).Oviparous animals show kinship effects in their micr obiomes, with hatc hlings fr om the same clutc h harboring a more similar cohort of microbes than unrelated animals (Yuan et al. 2015, Tr e v elline et al. 2018, Ambr osini et al. 2019 ).For many species, this inoculation is dependent on behavioral mechanisms associated with par ental car e .For example , se v er al bird species have been found to deposit skin, feather, preen oil, and fecal bacteria onto their eggs during incubation (Giraudeau et al. 2014, Martínez-García et al. 2015, van Veelen et al. 2018 ) and may influence the nest micr obial envir onment thr ough selection of plant materials with antimicrobial properties (Ruiz-Castellano et al. 2016 ).Some insect species will supply offspring with a specially pr oduced ca psule of obligate bacterial symbionts in the nesting en vironment (Hosoka wa et al. 2006 ), and some species of squid have a specialized organ for depositing bacteria into the jelly coat of their eggs (Nyholm 2020 ).
For egg-laying animals that do not pr ovide par ental car e (whic h includes over 99% of oviparous lizards and 97% of oviparous snakes; Richard 1987 ), inoculation with essential vertically transmitted micr obes m ust occur during egg de v elopment or oviposition.Microbes found within egg contents and internal egg surfaces can be traced to the maternal gut or r epr oductiv e micr obiome (Singh et al. 2014, van Veelen et al. 2018 ), indicating colonization during egg maturation.In chickens and rock pigeons, these microbes persist at least into the embryonic or hatchling stage (Lee et al. 2019, Dietz et al. 2020 ).Further, eggshells can be colonized with microbes from the cloaca during oviposition (van Veelen et al. 2018, Bunker et al. 2021, Li et al. 2022 ), and some bacterial species are able to penetrate eggshells (Gantois et al. 2009, Chen et al. 2019 ).These two potential pathways for maternal inheritance-through inoculation during egg development or penetration of the eggshell after oviposition-need further investigation, particularly as the maternal microbiome is not uniform thr oughout the digestiv e and r epr oductiv e tr act (Kohl et al. 2017, Bunker et al. 2022 ), and different microbial cohorts may be passed down from different maternal tissues.
Here , we in vestigate pathwa ys for maternal transmission of microbes in striped plateau lizards ( Sceloporus virgatus ), an oviparous phyrnosomatid lizard found in Mexico and parts of the southwestern USA.Sceloporus virgatus houses a specialized cloacal micr obiome, whic h is known to be tr ansferr ed to eggshell surfaces during oviposition, and to protect eggs from fungal infection during de v elopment (Bunker et al. 2021 ).Further, variation has been found in the microbiota of S. virgatus oviductal, cloacal, and intestinal tissue (Bunker et al. 2022 ), allowing for identification of the potential source of inherited microbes .T hese three maternal tissues are re presentati ve of the re producti ve microbiome (oviduct), the gut microbiome (intestine), and the junction of the two (cloaca).This system offers the opportunity to determine if maternal transmission is occurring in the lizards, and if so, to assess potential pathways of this transmission.

Sample collection
A total of 10 gravid female striped plateau lizards [ S. virgatus ; mean snout-to-vent length (SVL): 66.7 ± 0.73 mm (SE)] were collected by lasso from one contiguous population following r oughl y 2 km of creekbed near the Southwestern Research Station in Cochise County, Arizona, on 26-27 June 2022.Gravidity was readily determined by palpation.Lizards were k e pt in 23 cm × 15 cm cages sterilized with 70% ethanol prior to animal capture and had sterile water available ad libitum , but were not fed.S. virgatus females do not oviposit in captivity without induction.T hus , on 29 J une , after we observed females nesting in the field and w e determined b y palpation that eggs of ca ptiv e females wer e full y tur gid and r eady for oviposition, we induced the ca ptiv e females to oviposit via a 0.1-ml intr a peritoneal injection of oxytocin (Andr e ws and Rose 1994 ).Eac h female oviposited in individual cov er ed containers, lined with pa per to w el sanitized with 70% ethanol.Gener all y, oviposition began within ∼60 min of injection, and all females completed laying between 3.5 and 5.5 h of injection.We collected eggs as they were laid using sterile forceps, and weighed them (mean egg mass: 0.37 ± 0.002 g).
Clutc h size r anged fr om 11 to 15 eggs.For eac h clutc h, we destructiv el y sampled the first and last laid egg for contents and eggshell (described belo w), w e incubated tw o to three eggs in sterile vermiculite with no manipulation, and we treated the remaining eggs with a single species of fungal spores on Day 7 of incubation, as part of another study (see Table S1 , Supporting Information ).To acquire egg contents ( n = 20), we punctured each egg with sterile scissors and extruded the entire contents (including the embryonic disk, yolk, and all other extraembryonic material) into sterile 1.5 ml microcentrifuge tubes.Emptied eggshells ( n = 20) wer e stor ed in separate sterile microcentrifuge tubes.Efforts were made to k ee p eggshell and egg content samples distinct, but due to the nature of sampling there was still contact between eggshells and contents during collection.Incubated eggs ( n = 105) were placed in individual containers filled with sterile vermiculite dampened with 0.8 ml sterile water per gram; containers wer e cov er ed with par afilm and incubated at 30 • C until hatc hing.A sample of sterile v ermiculite, substr ate fr om the la ying surface , and a control swab of the r esearc hers' hands and general lab envir onment wer e taken as contr ols, to account for potential contamination.
On 30 J une , females were euthanized with two injections of buffered tricaine methanesulfonate, according to Conroy et al. ( 2009 ), follo w ed b y deca pitation.Tissue samples fr om the oviduct, cloaca, and intestine were taken with heat sterilized instruments.Oviduct and intestine wer e eac h sampled in two locations (oviduct: lo w er right and upper left oviduct; intestine: ∼2 mm above the cloaca and below the cecum), which were sequenced separ atel y and tr eated as r eplicates, as the comm unities wer e similar.We took a single, internal cloacal tissue sample from dir ectl y abov e the v ent.Substr ate fr om the dissection surface was taken as a control, to account for potential contamination.
There was a 100% hatch success rate for incubated eggs.Within 24 h of hatc hing, hatc hlings wer e r emov ed fr om their incubation cups and immediately weighed (mean hatchling body mass: 0.42 ± 0.003 g), measured (mean SVL: 24.0 ± 0.06 mm), and sacrificed by decapitation.We sampled the entire intestinal tr act, fr om just below the stomach to the cloaca, using a dissecting microscope and heat sterilized instruments, and wearing gloves sterilized with 70% ethanol.We did not subsample the intestine due to its small size (mean mass of hatchling tissue samples: 3.6 mg ± 0.19).Substr ate fr om the dissection surface, whic h was sterilized with 70% ethanol between eac h hatc hling, was taken as a control, to account for potential contamination.
All samples wer e stor ed at −80 • C until extraction.All methods were approved by the University of Puget Sound Institutional Animal Care and Use Committee (IACUC #PS21003).Fieldwork was conducted under Arizona Game and Fish Department License SP762402 and US Forest Service Special Use Permit DOU2223.

DN A extr action and Illumina libr ary prep
We extracted DNA from all tissue samples using the Qiagen DNEasy Blood and Tissue Kit (Qiagen, Inc), including the optional lysis buffer incubation for 30 min at 37 • C. Samples were incubated at 56 • C for 180 min while shaking at 500 RPM.Eggshells were rinsed with 200 μl sterile phosphate buffered saline (PBS) prior to extraction to remove any remaining vermiculite.We extracted DNA from eggshells similarly to tissue samples, with the addition of a bead beating step in which two sterile 2.8 mm ceramic beads were added to tubes prior to the second incubation and samples w ere v ortexed for 15 min at top speed.Eggshells were incubated at 56 • C for 90 min while shaking at 500 RPM.We extracted egg contents using the Qiagen Po w erSoil Kit, follo wing manufacturer pr otocols.All extr actions included an extraction blank as negative control, and PBS was added to the blank for relevant extractions.
Briefly, we pr epar ed Illumina libr aries via a two-step pol ymer ase c hain r eaction (PCR), in whic h PCR1 amplified the V4 region of the 16 s rRNA gene via 515F/806R primer pairs, and PCR2 added unique barcodes to each sample.Samples were pooled with varying volumes to qualitatively match DNA concentration, based on PCR band strength, and sent to the University of Idaho Genomics and Bioinformatics Core for clean up and sequencing on the Illumina MiSeq platform (v3, 2 × 300).More details can be found in Bunker et al. ( 2021Bunker et al. ( , 2022 ) ).A moc k comm unity was also amplified and included in sequencing as a positiv e contr ol and to inform data processing.

Raw data processing
We r eceiv ed sequences dem ultiplexed, with ada pters and primers r emov ed.All r aw data files have been uploaded to the NCBI Sequence Read Arc hiv e and can be accessed here: http://www.ncbi.nlm.nih.gov/bioproject/ 1023543 .Inspection of the mock community and quality scor es gener ated with FastQC (Andr e ws 2010 ) and a ggr egated with MultiQC (Ewels et al. 2016 ) were used to determine all parameters used in pr epar ation of r aw data.Samples wer e pr ocessed in R v 4.1.2using the D AD A2 pipeline (Callahan et al. 2016a ,b ).Samples were trimmed at 260 base pairs for forw ar d reads and 190 for reverse, and filtered with a maximum expected error of 2. Taxonomic classification of amplified sequence variants (ASVs) was performed through the assignTaxonomy function, using the Silva database (Quast et al. 2013 ), release 138.Potential contaminants were removed with the Decontam package (Davis et al. 2018 ), using the "pr e v alence" method with a threshold of 0.1.Control samples ( n = 31) included environmental controls from field collection, laying environment, and dissection surfaces, experimental contr ols, extr action blanks, and PCR negatives.We discarded any ASV that had fewer than 27 reads across all samples (based on analysis of mock communities) and all nontarget (nonbacterial) ASVs.Pr ocessed r eads wer e used to gener ate and optimize phylogenetic trees using the DECIPHER and Phangorn pac ka ges (Wright 2016, Schliep et al. 2017 ).Finally, read numbers were log transformed to account for differences in read depth, as r ar efaction curv es indicate sufficient div ersity was ca ptur ed for all samples, and thus r ar efaction was not necessary ( Figure S1 , Supporting Information ).We excluded samples which had fewer than 500 reads after processing from the analysis ( n = 9).Final sample sizes are in Table 1 .All figur es wer e made using the gg-plot2 pac ka ge (Wic kham 2017 ).

Sta tistical anal ysis
All anal yses wer e performed in R (v ersion 4.1.2)(R Cor e Team 2020 ).Phyloseq (McMurdie andHolmes 2013 et al. 2013 ) was used to organize data of different types (i.e .ASV counts , taxonomy, and metadata).Shannon diversity and richness values were generated using the "estimate_ric hness" function fr om phyloseq, and Faith's phylogenetic diversity (PD) was estimated using the described phylogenetic trees and the Picante package (Kembel et al. 2010 ).
Principal coordinates analysis (PCoA) plots wer e gener ated using the "ordinate" function, and weighted and unweighted UniFrac distance matrices were generated using the function "UniFrac," both from the phyloseq package.
To investigate the potential for maternal microbiota transmission, we assessed the impact of maternal identification on hatchling intestinal microbiota.All successfully hatched offspring were included in these analyses as the fungal treatment did not have an effect on offspring microbiota ( Figures S2 and S3 , Supporting Information ), and treatments were equally represented within a clutch ( Table S1 , Supporting Information ).Effect of maternal ID on alpha diversity was tested with ANOVAs, controlling for extr acted hatc hling tissue mass in milligrams, after determining that the models met assumptions of normality and dispersion.Onl y ric hness was log tr ansformed to meet these assumptions.Ov er all effect of clutch on community membership and composi-tion (also controlling for sample mass) was tested with permutational ANOVAs (permANOVA) using the "adonis2" function from the v egan pac ka ge (Oksanen et al. 2019 ).Community dispersion was tested with the "betadisper" function, also from vegan.
To examine pathways of transmission from maternal tissues (oviduct, cloaca, and intestine) to all three offspring sample types (egg contents , eggshells , and intestine), the remaining analyses only included intestinal samples from "control" hatchlings, which were incubated in a sterile environment with no fungal treatment ( n = 2-3 per clutch; Table S1 , Supporting Information ).First, we compared alpha and beta diversity metrics across all maternal and offspring sample types.Alpha diversity was compared using ANOVAs with maternal ID as a block, and again only richness was log transformed to meet assumptions.Tuk e y HSD tests were used for post hoc pairwise comparisons.Ov er all comm unity composition and structure were compared with permANOVAs and beta dispersion, as described abo ve .Post hoc pairwise comparisons were performed with the pairwiseAdonis pac ka ge (Martinez 2017 ).Second, w e used Sour ceTr ac ker (Knights et al. 2011 ) to identify the likely maternal source tissue for offspring microbes, with all maternal tissues defined as "source" and all offspring sample types as "sink."SourceTr ac ker identifies the likely source of ASV's but does not assess community structure .T hus , third, we compared community composition of individual offspring sample types to all maternal tissues.PCoA plots were generated using weighted and unw eighted UniF r ac distance matrices.Eac h plot was gr ouped into clusters of samples with k-means clustering.The number of clusters for each plot was determined based on elbow plots and av er a ge silhouette width v alues.Clusters wer e gener ated with the "pam" function from the cluster pac ka ge (Maec hler et al. 2022 ).
Finally, to assess variation in microbe provisioning within a clutc h, we compar ed the egg content and eggshell micr obiota of the first and last egg laid by each female .T he influence of lay order on alpha diversity was examined using paired t -tests; all metrics except eggshell Shannon diversity were log transformed to meet test assumptions .T he influence of la y or der on beta diversity w as examined using beta dispersion tests and permANOVAs, controlling for maternal ID, as described abo ve .
Data analysis files have been submitted to the Dryad data repository, and can be accessed here: https:// datadryad.org/stash/ share/MGM1O8QrDcWxRdjIoUaudfk-e _ GTK0G _ GPHQhTsmzpA

Offspring sample community compositions
For all three offspring sample types, the most abundant families were Enterobacteriaceae and Yersinaceae (Phylum Proteobacteria; Figure S4 , Supporting Information ).Members of the Enterobacteriaceae family made up 45.4 ± 7.1% of the egg contents community, 63.8 ± 7.6% of the eggshell community, and 53.8 ± 3.4% of the hatchling intestine community, on average (Fig. 1 A).Members of the Yersinaceae family made up 31.8 ± 7.4% of the egg contents community, 21.1 ± 7.2% of the eggshell community, and 22.2 ± 3.2% of the hatchling intestine community, on a verage .T he most common and abundant member of Enterobacteriaceae was Klebsiella , while all Yersinaceae ASVs were Serratia or unidentified at the genus le v el (Fig. 1 B).No other famil y made up mor e than 5% of the av er a ge egg contents community.Eggshells and hatchling tissues each had one additional family accounting for ∼5% of the community on av er a ge ( Pseudomonadaceaea and an unknown Enter obacter ales, r espectiv el y), with the remainder of the community for all

Ma ternal tr ansmission
We used intestinal tissues from all hatched offspring tissue to identify effects of maternal ID on hatchling microbiota.We found that all three alpha diversity measures of hatchling intestine tissue (Shannon diversity index, richness, and PD) var-   Figur e 5. T he % of each sample type assigned to each cluster (most closely grouped samples based on cluster analysis) for weighted UniFrac distance (A), and unw eighted UniF rac distance (B).Colors r epr esent eac h cluster, and height of each color r epr esents the % of each sample type assigned to that group.ied depending on maternal ID (F 9,89 ≥ 3.04, P ≤ .018;Table S2 , Supporting Information ).As well, both beta diversity metrics of hatchling intestine tissue varied with maternal ID (weighted and unw eighted UniF rac; P = .001for both; Table S3 and Figure S5 , Supporting Information ), with maternal ID accounting for 21% of variation in the hatchling microbiota based on weighted UniFrac distance, and 16% based on unw eighted UniF rac distance.Samples were also dispersed differ entl y between clutc hes based on unw eighted UniF rac distance (F 9,89 = 2.51, P = .013,Table S3 , Supporting Information ), although not weighted UniFrac (F 9,89 = 1.37,P = .212,Table S3 , Supporting Information ).

Pathways of transmission
To examine potential pathways of microbe transmission, we compared egg contents , eggshells , and control hatchling intestine tissue (i.e.animals hatched from eggs which did not receive a fungal inoculation) to three maternal tissue types: oviduct, cloaca, and intestine.ANOVAs of all alpha div ersity measur es indicate an ov er all differ ence in div ersity between all maternal and off- spring sample types (F 5,93 ≥ 2.31, P ≤ .022;Table S4 , Supporting Information ; Fig. 2 ).Based on post hoc tests, alpha diversity of egg contents , eggshells , and hatchling tissues were similar to the maternal cloaca and lo w er than the maternal intestinal tissue, in all metrics ( Table S5 , Supporting Information ).Relative to the maternal oviductal tissue, egg contents had significantly lo w er diversity in all measures, eggshells had statistically similar diversity in all measur es, and hatc hling tissues had significantl y lo w er diversity for Shannon and PD, but not for richness ( Table S5 , Supporting Information ).PermANOVAs and pairwise permANOVAs indicate differences between the community structure of all sample types ( P ≤ .010;Tables S6 and S7 , Supporting Information ), although patterns of ov erla p wer e still observ ed between gr oups on PCoA plots (Fig. 3 ), whic h wer e explor ed further with clustering anal yses (see below).
The SourceTr ac ker anal ysis found that an av er a ge of 35.6% of ASVs in the egg contents w ere sour ced from the maternal oviduct and 34.4% of ASVs were sourced from the maternal cloaca (Fig. 4 , Supplemental File 2 ).In contrast, the majority of ASVs found on eggshells and in hatchling tissue were sourced from the ma-ternal cloaca on av er a ge (58.4% and 64.3%, r espectiv el y), with most of the remaining ASVs coming from the oviduct (25.6% of ASVs on eggshells and 20.7% of ASVs in hatchling tissue) (Fig. 4 , Supplemental File 2 ).For all offspring tissue types, a r elativ el y lo w per centage of ASVs w ere sour ced from the maternal intestine: 0.6% in egg contents, 1.0% on eggshells, and 1.5% in hatchling tissue (Fig. 4 , Supplemental File 2 ).The source of the remaining taxa was unidentified.
While the SourceTr ac ker anal ysis identifies the likel y maternal source of offspring ASVs, it does not assess the community structur e of eac h offspring sample type r elativ e to maternal sample types .T hus , we used cluster analysis based on the PCoA plots to assess whether the composition of offspring samples tend to cluster with a given maternal sample type.Using k-means clustering, samples wer e gr ouped into thr ee clusters, with eac h cluster primarily associated with one of the maternal tissue types (Fig. 5 ).When using weighted UniFrac distance, Cluster 1 contained 90% of the maternal intestine tissues, only 22% of maternal oviductal and 10% of maternal cloacal tissue samples, and only a single eggshell sample, with no egg contents or hatchling tissues.Clus- ter 2 included 61% of the oviductal samples and 10% of the cloacal samples, but none of the maternal intestinal samples.It also included the majority of the egg contents (56%), 48% of hatchling intestine samples, and 26% of eggshells.Cluster 3 contained 80% of the maternal cloacal samples, 16% of the maternal oviduct samples, and the remaining 10% of maternal intestine samples, as well as 44% of egg contents, 52% of hatchling intestines, and 68% of eggshells (Fig. 5 A).Using unweighted UniFr ac, a gain, Cluster 1 contained most of the maternal intestine samples (90%), Cluster 2 contained most of the maternal oviduct samples (83%), and Cluster 3 contained most of the maternal cloacal samples (80%).All three offspring sample types predominantly fit into Cluster 2: 78% of egg contents, 73% of eggshells, and 87% of hatchling intestine, with nearly all other samples fitting into Cluster 3 and a single eggshell sample fitting into Cluster 1 (Fig. 5 B).

Within-clutch v aria tion in tr ansmission
We found that contents of the first egg had significantly lo w er alpha diversity than did that of the last egg for both Shannon Diversity and Richness ( t 7 ≤ −2.41, P ≤ .047,Fig. 6 ), and there was a similar trend for PD ( t 7 = −2.08,P = .077;Table S8 , Supporting Information ; Fig. 6 ).There was no variation in community structure or membership ( Table S9 , Supporting Information ; Fig. 7 A and B).The shell of the first egg laid in eac h clutc h had significantly lo w er diversity than the shell of the final egg in all three alpha diversity measures ( t 8 ≤ −2.78, P ≤ .024;Table S8 , Supporting Information ; Fig. 6 ).Ov er all comm unity structure (w eighted UniF rac, P = .011,Fig. 7 A) and membership (unw eighted UniF rac, P = .043,Fig. 7 B) differed between the first and last eggshells, but dispersion did not ( Table S9 , Supporting Information ).

Discussion
We found that maternal ID impacted the microbiota of hatchling tissue, which supports the hypothesis that maternal transmission is occurring in S. vir gatus .T he micr obes ar e tr ansferr ed primaril y fr om the cloaca and oviduct during egg de v elopment and oviposition, with very little impact from intestinal microbes.Both egg contents immediately after oviposition and offspring tissues immediatel y after hatc hing most closel y r esemble the oviductal community.This indicates that microbes may colonize the egg during de v elopment, possibl y befor e the shell is de v eloped, and these microbes persist through development into the hatchling intestine .T his has been observed in chickens, in which the early egg white microbiome as well as the embryonic gut most closely resembled the oviductal community (Lee et al. 2019 ).Similarly, the gut microbiota of Amazon river turtle hatchlings is dependent on the early egg microbiota, although it was lar gel y acquir ed fr om envir onmental micr obes r ather than maternal tissues (Carr anco et al. 2022 ).
There is also a significant influence of cloacal micr obes, whic h make up the majority of the taxa in all offspring communities, despite the ov er all structur e of the communities being most similar to the o viduct.T hese cloacal microbes appear to be primarily tr ansferr ed via the outer eggshell.This shell microbiome is known to protect eggs from fungal fouling during incubation and has also been associated with increased hatchling fitness (Bunker et al. 2021 ).The major taxa in both the maternal cloaca and eggshells belong to the Yersinaceae and Enterobacteriaceae families, which have been identified as important functional taxa in adult cloacal microbiomes, as they provide antifungal protection to eggs during de v elopment in the soil (Kalbe et al. 1996, Dhar Purkayastha et al. 2018, Bunker et al. 2021 ).This indicates some selection for these microbes, and could explain the pr efer ential v ertical tr ans-mission observ ed her e.Some bacterial taxa ar e able to penetrate eggshells (Gantois et al. 2009 ), so it is possible that these microbes are deposited on the shell only during oviposition and establish in the embryo during incubation.It has also been hypothesized that microbes can ascend to the re producti ve tract from the vagina (in humans; Hansen et al. 2014 ) or gut (in c hic kens; Shterzer et al. 2020 ), raising the possibility that the "cloacal" microbes also establish in the internal egg environment prior to shelling in the oviduct.
Because offspring communities have a high percentage of cloacal taxa but r elativ el y low similarity with the cloacal community structur e, ther e m ust be physiological or envir onmental pr operties , which fa vor the growth of certain taxa in eggs and hatchlings as they de v elop.For example, egg albumen is known to have antimicr obial pr operties (Shawk e y et al. 2008 ), which may select for bacteria whic h ar e r esistant to the antimicrobial peptides present in the egg.Further, the cloaca may be a more aerobic region than the upper areas of the oviduct and gut, which could impact how well the associated microbes persist in the egg environment during incubation and in the different regions of the hatchling gut or r epr oductiv e tr acts (Shawk e y et al. 2008, Vide v all et al. 2018, Berlow et al. 2020 ).This could also account for the ov er all v ariation between sample types, as only a subset of the maternal bacteria survived in the egg environment.
The variation between first eggshell and last eggshell communities also indicates that microbes are not evenly provisioned to offspring.Given that the same patterns were not observed in the egg contents, variation in eggshell microbiota likely occurred after the eggshell formed.Eggs that de v elop higher in the oviduct, and thus have to pass through the entire length of the oviduct during oviposition would likely be exposed to a greater diversity of microbes, impacting the eggshell microbiota but not the egg contents at the time of sampling.van Veelen et al. ( 2018 ) found that the eggshell microbiome of first and second eggs in two lark species varied in the relative abundance of particular taxa, but these micr obes wer e sourced to the nest envir onment r ather than the maternal tissues.Mor e r esearc h acr oss ovipar ous taxa is needed to assess the effect of lay order on eggshell microbiota and the sources of those microbes.
Unlike the cloacal and oviductal microbes, we found little evidence that maternal intestinal microbes colonize eggs or hatchlings.Studies in c hic kens hav e found that gut micr obes ar e tr ansmitted to eggshells (Ding et al. 2017, Shterzer et al. 2020 ), although in S. virgatu s the distinct differentiation of the cloacal microbiome and inability of fecal microbes to establish at the cloaca (Bunker et al. 2022 ) make it less likely that eggs would come into direct contact with upper gut microbes.Ho w ever, the lack of intestinal micr obes was sur prising in the hatc hling tissue samples as the entir e gut r egion was sampled.This indicates that v ariation between gut and r epr oductiv e tissues de v elops as the animals age, and that the adult intestinal microbes are likely acquired from diet or the en vironment.T here is evidence in humans (Koenig et al. 2010 ), other mammals (Wang et al. 2019), bir ds (Hir d et al. 2014, Taylor et al. 2019, Vide v all et al. 2019, Hernandez et al. 2021 ), and other reptiles (Yuan et al. 2015 ) that the micr obiome under goes lar ge structur al c hanges both immediatel y after birth/hatc hing and as animals age from juveniles to adults, which may also account for differentiation of gut regions .T his change could also be instigated when the animals begin for a ging, offering an inter esting ar ea of investigation for future research.
This study was conducted in a controlled, sterilized environment, and does not account for the potential impact of nest micr obes, whic h ar e known to be integral to de v elopment of the neonate microbiome in other species (van Veelen et al. 2018 , Campos-Cerda andBohannan 2020 ).Although the lack of parental care means that S. virgatus nests are not manipulated in the way that bird and other reptile nests are, the nest microbiome has still been shown to influence offspring microbes in other reptiles that also lack parental care (Carranco et al. 2022, Li et al. 2022 ).We cannot rule out the possibility that soil micr obes penetr ate the shell during incubation to colonize the embryo, leading to a different hatc hling micr obiota in wild hatc hlings than the one r ecov er ed in this stud y.Ad ditionally, we noted the presence of the incubation medium (vermiculite) in the hatchling intestines upon dissection, indicating the possibility that hatchlings ingest soil as they leave the nest.This could serve as an initial inoculation with soil microbes, altering the wild hatchling microbiota and accounting for micr obial v ariation found in other lizard species based on habitat or geogr a phic location (Baldo et al. 2018, Aleman y et al. 2022, Bunker and Weiss 2022 ).
Finall y, ther e ar e a lar ge percenta ge of micr obes pr esent in hatc hlings that wer e not found in an y of the maternal tissues we sampled, and it is unclear where those taxa are originating from.It is possible that the taxa ar e pr esent in low le v els in the maternal tissues, but wer e onl y detectable in the r elativ el y lo w er bioload environment of the offspring samples.Some of these taxa may originate from tissues which were not sampled in this study, particularl y the ov aries, whic h hav e been hypothesized as a source for gec k o egg micr obes (Singh et al. 2014 ) or the upper intestine, which is known to be differentiated from the lo w er intestine and o viduct in S. vir gatus (Bunker et al. 2022 ).Another possibility is that these microbes could originate from the paternal ejaculate microbiome; while less well studied than the female r epr oductiv e micr obiome, micr obes hav e been isolated from sperm and in some cases can impact r epr oductiv e success (Ro w e et al. 2020 ).
Ov er all, these r esults establish that maternal tr ansmission of microbes is occurring in S. virgatus , and much of that transfer occurs prior to oviposition, within the oviduct.Mor e r esearc h m ust be done to see how this transfer is complicated by soil microbes, how the microbiome differentiates in different tissues over time, and the impact of paternal microbes .T hese findings can serve as a baseline to identify potential pathways of maternal transmission in other oviparous animals, and shift the primary focus from the gut to the r epr oductiv e micr obiome.

Figure 1 .
Figure 1.Av er a ge comm unities of all sample types.Mean r elativ e abundances of top 10 most abundant Families (A), and Gener a (B) acr oss sample types, with the remaining taxa grouped into the "Other" category.Colors represent different taxa, and the height of each bar represents the average % of total reads assigned to each taxa, in each sample type.

Figure 2 .
Figure 2. Shannon Diversity index values , Richness , and Faith's PD index values for all sample types .P oints represent individual diversity value for each sample, bo xes re present median and quartiles of group diversity.The shape of the points indicates maternal (squares) or offspring (circles) samples.

Figure 3 .
Figure 3. PCoA plots based on weighted (A) and unweighted (B) UniFrac distances, comparing three offspring sample types and three maternal tissue sample types.Colors of points and lines r epr esent differ ent sample types, and the sha pe of points indicates maternal (squar es) or offspring (circles) samples.Filled points r epr esent individual samples, and larger open points represent centroids from each group.

Figure 4 .
Figure 4. Av er a ge % of ASVs sourced from each maternal tissue type or unknown source (r epr esented by colored bars) for each offspring sample type.

Figure 6 .
Figure 6.Shannon Diversity index values , Richness , and Faith's PD index values for egg contents and eggshells of first and last laid egg in each clutch.Colors r epr esent sample types and sha pes r epr esent egg lay order.

Figure 7 .
Figure 7. PCoA plots based on weighted (A) and unweighted (B) UniFrac distances, comparing egg contents and eggshells of first and last laid egg in eac h clutc h.Colors r epr esent sample types and sha pes r epr esent egg lay order.Closed sha pes r epr esent individual samples, and lar ger open sha pes r epr esent centr oids fr om eac h gr oup.

Table 1 .
Final sample size used in analyses for each sample type.
*Eggs not treated with fungus on Day 7 of incubation.