A symbiotic physical niche in Drosophila melanogaster regulates stable association of a multi
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A symbiotic physical niche in Drosophila melanogaster regulates stable association of a multi

Dec 15, 2023

Nature Communications volume 14, Article number: 1557 (2023) Cite this article

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The gut is continuously invaded by diverse bacteria from the diet and the environment, yet microbiome composition is relatively stable over time for host species ranging from mammals to insects, suggesting host-specific factors may selectively maintain key species of bacteria. To investigate host specificity, we used gnotobiotic Drosophila, microbial pulse-chase protocols, and microscopy to investigate the stability of different strains of bacteria in the fly gut. We show that a host-constructed physical niche in the foregut selectively binds bacteria with strain-level specificity, stabilizing their colonization. Primary colonizers saturate the niche and exclude secondary colonizers of the same strain, but initial colonization by Lactobacillus species physically remodels the niche through production of a glycan-rich secretion to favor secondary colonization by unrelated commensals in the Acetobacter genus. Our results provide a mechanistic framework for understanding the establishment and stability of a multi-species intestinal microbiome.

Host health is affected by the composition of the gut microbiome, specifically which species and strains of bacteria occupy the gut1,2,3,4,5. The microbiome is established and maintained in the face of daily fluctuations in diet, invasion by pathogens6, and disruptions by antibiotics7. Many gut resident bacteria localize to specific regions of the gut that correspond to chemical environments matching the specific species’s metabolism8. Certain probiotics, namely Lactobacillus species, additionally make physical attachments with host mucus, stabilizing their colonization9,10. With strain-level diversity in the hundreds to thousands11, it remains enigmatic how a host can select and maintain a specific set of strains. One hypothesis is that long-term maintenance of diet and lifestyle habits reinforce microbiome stability12,13,14,15,16, while another, non-exclusive hypothesis is that the host constructs physical niches in the gut that acquire and sequester specific symbiotic bacteria17,18,19,20,21,22.

The microbiome of the fruit fly, Drosophila melanogaster, has been studied for over a century and is relatively simple in its composition compared to the mammalian gut microbiome23, yet how fly gut microbiome assembly is regulated remains unclear. Similar to mammalian colonic crypts, the fly gut is microaerobic and colonized by bacteria from the Lactobacillales class and Proteobacteria phylum22,24,25,26. Flies can easily be reared germ-free and then associated with defined bacterial strains, providing a high level of biological control27. Furthermore, the fly gut microbiome has low diversity, with ~5 species of stable colonizers from two primary groups: the genera Lactobacillus (phylum Firmicutes), which was recently split into Lactiplantibacillus and Levilactibacillus, and Acetobacter (class α-Proteobacteria)26,28. These species are easily cultured, genetically tractable27, and they affect fly lifespan, fecundity, and development29,30,31,32,33,34,35. While colonization of the fly gut has long been argued to be non-specifically regulated by host filtering mechanisms, including feeding preferences, immunity, and digestion, recent evidence suggests flies may also selectively acquire Lactobacillus and Acetobacter strains in the wild24,36, and these may provide flies with nutrition during the larval phase37.

Here, we discover a physical niche within the adult Drosophila foregut that is specifically colonized by wild strains of Lactobacillus and Acetobacter. We characterize the spatial specificity of the niche, the bacterial strain specificity for colonization, and the stability of colonization. We measure priority effects that regulate the order in which the bacterial species colonize. Finally, we measure the response of the niche to bacterial colonizers including physical changes and glycosylation of the extracellular matrix.

In previous work, we surveyed a range of bacterial strains that were associated with either lab or wild-caught D. melanogaster24, identifying a subset of bacterial strains that efficiently colonize the gut of lab flies. To investigate whether commensal bacteria form stable associations with the fly gut in a manner consistent with the existence of a niche, we exposed flies to a quantified inoculum of bacterial cells labeled with a fluorescent protein (Fig. S1A–G). Following inoculation, flies were transferred to germ-free food daily for 3 d followed by an additional transfer to a new germ-free vial for 3 h to allow transient bacteria to clear from the gut (“Methods”, Fig. S1). Clearing prior to analysis reduced the total number of gut bacteria and the spatial variation in bacterial location (Fig. S1H–J). These experiments revealed that a strain of Lactiplantibacillus plantarum (Lp) isolated from a wild-caught fly (LpWF) persists exclusively in the D. melanogaster foregut (Figs. 1A–D, S1I,J), including the proventriculus (a luminal region connecting the esophagus with the anterior midgut38), the crop (a sack-like appendage), and the crop duct that connects the crop to the proventriculus. Bacteria associated with longitudinal furrows lining the surface of the proventriculus inner lumen, the crop duct, and the base of the crop (Fig. 1C-D, S1J). Similar spatial patterns of colonization occurred in adult mated females, virgin females, and adult males (Fig. S2A–J), indicating that sex-specific differences do not affect the phenotype. No colonization was observed in larvae post-clearing, indicating that the phenotype is specific to the adult foregut (Fig. S2K–M). We focus on adult mated females throughout because they have been shown to exhibit low fly-to-fly variation in gut phenotypes39,40,41.

A Colonization assay schematic with initial dosing on day 0 and serial transfers to sterile food daily for 3 days before analysis. B Gut diagram. C Microscopy of LpWF-mCherry colonization in a whole gut after clearing transient cells shows a specific colonization zone in the foregut. Shown is a maximum-intensity z-projection. D The proventriculus is a major site of LpWF colonization. E Ai colonization is also specific to the proventriculus lumen and crop duct (see also Fig. S2). F CFU densities from regions dissected in B. n = 23 individual guts/region from three biological replicates. Columns represent means. Error bars are S.D. G Microsurgery was carried out to remove the crop. H LpWF colonizes the foregut of flies with the crop removed (n = 15/15). I TEM cross-section of proventriculus inner lumen. Representative image of n = 3 biological replicates. J Detail of (I). Scale bars are defined in the figure panels. Source data are provided as a Source Data file.

Similar to LpWF, a strain of Acetobacter indonesiensis colonized the same foregut regions (Figs. 1E, S2, S3), indicating that the two major groups of fly gut bacteria have the same spatial specificity in the foregut. By contrast, flies colonized with Lp from laboratory flies (LpLF) (Fig. S1K, L) or the LpWCFS1 strain isolated from humans (Figs. S1M, S2A–J) exhibited much lower levels of colonization. No Lp strains were found at substantial abundance in the midgut or other regions of the fly after clearing transient bacteria. Consistent with microscopy, live bacterial density was greatest in the proventriculus, followed by the crop, and was lowest in the midgut and hindgut (Figs. 1F, S1N). We further validated that LpWF maintains stable colonization in the absence of ingestion of new bacterial cells over 5 d during which non-adherent bacteria were flushed from the gut by fastidiously maintaining sterility of the food using a CAFÉ feeder24 (Fig. S4A, B).

A bacterial population in the foregut with the observed spatial localization might be maintained by proliferation and constant re-seeding from the crop, in which case flies without crops could not be stably colonized. We conducted microsurgery to remove the crop from germ-free flies (Fig. 1G, “Methods”), inoculated them with LpWF 5 d post-surgery, and then dissected and imaged the gut 5 d post inoculation (dpi). Surgical success was validated and the remaining portion of the crop duct had a melanized scar at the surgery site (Fig. 1H). All cropless flies were stably colonized by LpWF (n = 15/15), with a high density of bacteria in the proventriculus inner lumen as in flies with an intact crop (Fig. 1C). We observed similar Ai colonization following cropectomy (n = 14/14 colonized, Fig. S3E, F). Examining these regions further, transmission electron microscopy (TEM) of the proventriculus lumen revealed a consistent tissue geometry (Fig. 1I, J), with densely packed bacterial cells longitudinally oriented in elongated furrows formed by host cell bodies making up an average of 11 ridges per cross section (Fig. S4C).

Thus, the crop is not required for stable foregut colonization by LpWF or Ai, suggesting that the ability to specifically bind to the proventriculus and crop duct is key to stable bacterial association.

A niche would be expected to result in strong bacterial association based on specific binding sites, such that the associated bacterial population size would saturate at a well-defined value. Moreover, cells already bound to the proventriculus would be expected to promote population stability and prevent later-arriving bacteria from colonizing. To test these hypotheses, we colonized germ-free flies with a range of doses of LpWF-mCherry and measured the abundance over time. As predicted, over a wide range of initial inoculum sizes, the associated bacterial population saturated at ~104 CFUs/fly (Fig. 2A). Furthermore, when the inoculum size was below that saturation level, the population of bacteria in the proventriculus increased gradually and plateaued within 5 d. Growth measurements in live flies24 demonstrated that the plateau was reached by growth of the initially bound population rather than ingestion of additional cells. By contrast, when an excess of bacteria was supplied initially, the population decreased to the same plateau value within 1 d (Fig. 2A), indicating that the niche has a finite and fixed carrying capacity. Similar dynamics were observed for Ai with ~103 cells at the saturated density (Fig. S3G).

A Saturation occurs over a colonization time course of germ-free flies by LpWF. Data points are mean of log10(CFUs) in n ≥ 48 flies/data point. Error bars represent 1 s.e.m. Inset: 20-day time course after inoculation with 106 CFUs (data from24). B Bacterial pulse-chase experimental design: flies were first pre-colonized with LpWF-mCherry, then fed an excess of unlabeled LpWF (blue) daily on fresh food. C Acterial cell turnover quantified by pulse-chase time course of Lp-mCherry-pre-colonized flies continuously fed unlabeled LpWF or Ai-GFP-pre-colonized flies continuously fed unlabeled Ai. Data points are mean of log10(CFUs) in n ≥ 34 flies/data point. Error bars represent 1 s.e.m. D Colonization efficiency quantified by dose response to colonization of individual flies. CFUs were measured at 3 dpi of the second colonizer. n = 24 flies/dose, error bars represent 1 standard error of the proportion. Limit of detection: 50 CFUs. E Spatial structure of colonization dynamics in the proventriculus for a fly pre-colonized with LpWF-mCherry (red) invaded by LpWF-GFP and imaged 1 h post inoculation (hpi). F Optical x,z-slice. Source data are provided as a Source Data file.

To investigate the stability of bacterial colonization in the proventriculus, we performed a pulse-chase experiment in which we challenged LpWF-mCherry-pre-colonized flies with unlabeled LpWF fed in excess over the course of 10 d (Fig. 2B). LpWF-mCherry levels in the gut decreased by >90% over the first 5 d, from ~104 to ~103 CFUs/fly, and then remained at ~103 CFUs/fly for the following 5 d (Fig. 2C), indicating a small, bound population with little turnover and a larger associated population with a half-life of 2.5 d (95% confidence interval (c.i.) 1.6–4.3 d). By contrast, LpWCFS1, a weakly-colonizing human isolate of L. plantarum, was quickly flushed from the gut (Fig. 2C). Similar dynamics were observed in Ai (Fig. 2C, S3H) with a half-life of 2.5 d (95% c.i. 1.3–6.5 d), indicating that the niche has equivalent kinetics for both bacterial species.

Initial binding to the niche is a key step in the establishment of a new bacterial population prior to filling the niche. Establishment is dose-dependent24, and our finding that the final abundance of late colonizers is lower than that of initial colonizers (Fig. S4D) suggested that the presence of prior colonizers would shift the dose-response curve. To quantify such priority effects, we fed a range of doses of LpWF-mCherry to individual LpWF-pre-colonized flies and measured the percentage that were colonized by LpWF-mCherry 3 d later. Consistent with our hypothesis, pre-colonized flies were less likely than germ-free flies to become colonized by an equal dose of LpWF-mCherry: ~103 LpWF-mCherry CFUs were required for 50% of flies to be colonized, while 100% of germ-free flies ended up colonized by doses as low as 102 CFUs (Fig. 2D). These findings demonstrate that the proventricular niche for LpWF, when occupied, strongly resists colonization by later doses of the same strain.

The relationship between the probability of establishment and the final abundance of successful colonizing bacteria suggests that the availability of open habitats regulates the chance of invasion. We formalized assumptions of this hypothesis by building an integrated theory of initial colonization24 and niche saturation42 that predicts the likelihood of colonization, \(P\left({N}_{0}\right)\), of an invading species inoculated at a dose of \({N}_{0}\) as a function of the final abundance of the invading species, \({A\left (\right. N}_{0}\left )\right.\):

where \(p\) is the colonization probability of an individual bacterial cell and \(k\) is the subpopulation size attained in a single successful colonization event (Fig. S5A, B). This model allowed us to estimate the scale at which the population is structured based on colonization probabilities and total bacterial abundances. For LpWF, Eq. 1 estimates a subpopulation size of \(k\)=600 cells (Fig. S5C), which is roughly the number of cells contained in an individual furrow.

To test whether the later dose of LpWF-mCherry was spatially excluded by resident LpWF, we constructed a GFP-expressing strain of LpWF and fed it to flies pre-colonized with LpWF-mCherry. We imaged whole fixed guts 1 h post inoculation (hpi) to capture LpWF-GFP cells before they passed out of the fly (Fig. 2E). In the proventriculus, the invading LpWF-GFP were localized along the central axis of the inner lumen, separated from the lumen wall by a layer of resident LpWF-mCherry (Fig. 2E, F) that was up to 10 µm thick. The posterior proventriculus furrows were densely packed with LpWF-mCherry, while LpWF-GFP was largely absent from furrows, suggesting that these furrows are the sites of stable colonization. We confirmed that the fluorophores are not responsible for the differential colonization by feeding LpWF-mCherry to flies pre-colonized by unlabeled LpWF and quantifying the mCherry signal along the gut at 1 hpi and 24 hpi. At 24 hpi with a dose of ~104 CFUs, flies pre-colonized by LpWF showed almost undetectable mCherry by microscopy (Fig. S4E–H). These results provide further support that the niche for LpWF is in the proventricular furrows. Unlike during initial colonization, in which bacteria rapidly enter and colonize the furrows, prior colonizers prevented subsequent colonization, suggesting that there are a limited number of binding sites in the furrows for LpWF cells and that these sites are saturated by prior colonization. Consistent with this logic that niche priority is spatially determined, in the cases when LpWF-GFP did show colonization (n = 5), GFP-labeled cells were co-localized with each other along a furrow rather than being evenly mixed with mCherry (Fig. S4I).

Interspecies interactions can have major impacts on ecosystem colonization through priority effects that include competitive exclusion and facilitation43,44,45,46,47. Because Ai and LpWF colonize the same general location of the gut (Figs. 1C–F, S3A–D) and each strain excludes itself (Figs. 2D, 3A), we expected that they would exclude each other. To test this hypothesis, we measured each species’s abundance and growth rate during co-colonization. To our surprise, both were unaffected (Figs. 3B, C, S6), demonstrating that the species independently saturate the niche. We also performed a dose-response assay to determine whether interactions affect establishment of new colonizers. By contrast to Ai’s self-exclusion, LpWF pre-colonization facilitated Ai colonization (Fig. 3A), while LpWF colonization was unaffected by the presence of Ai (Fig. S6A–C). A. pasteurianus, a phylogenetically distinct species of Acetobacter48 that is common in D. melanogaster49, was also facilitated by LpWF (Fig. S6D). Heat-killed LpWF did not facilitate Ai colonization, indicating live LpWF cells are necessary to facilitate Ai colonization (Fig. S7A, B).

A Strain interactions influence colonization efficiency, as seen by dose-response curves for Ai fed to germ-free flies (open green circles), Ai-pre-colonized flies (filled yellow circles), or Lp-pre-colonized flies (black-filled green squares). Z-test of differences in proportion versus Ai into germ-free flies: dose 102.3 CFUs/fly, p = 8.1×10−4; dose 103.7 CFUs/fly: p = 4.8 × 10−9; dose 105 CFUs/fly: p = 8.7 × 10−6. n ≥ 12 flies/data point. Error bars represent 1 standard error of the proportion. B Ai abundance at 5 dpi does not differ between flies mono-colonized with Ai versus pre-colonized with LpWF then fed Ai. n ≥ 65 flies/treatment; two-tailed unpaired t-test, p = 0.38; ns indicates not significant. C LpWF abundance 5 dpi does not differ between flies mono-colonized with LpWF versus pre-colonized with Ai then fed LpWF. n ≥ 53 flies/treatment; two-tailed unpaired t-test; p = 0.06; ns indicates not significant. B, C: Center of box is median; box encloses 25th to 75th percentiles; whiskers indicate minimum and maximum. D Confocal microscopy of Lp and Ai co-colonization. Ai (green) and LpWF (red) occupied the same regions of the foregut 1 dpi. Scale bar: 100 µm. E, F x,z-section of Ai and LpWF sectors. G TEM cross-section of Ai and LpWF co-colonizing the anterior proventriculus. Scale bar: 5 µm. H Detail of (G) with LpWF and Ai cells pseudocolored. Scale bar: 2 µm. Source data are provided as a Source Data file.

Fluorescence microscopy of guts co-colonized by LpWF-mCherry and Ai-GFP showed that Ai and LpWF co-colonized the same foregut regions (Fig. 3D), with distinct sectors of each species (Fig. 3E–H, Supplementary Movie 1). Thus, LpWF and Ai do not physically exclude one another; instead, the tissue accommodates both strains.

To examine the coexistence of overlapping Ai and LpWF populations in a physically confined space, we imaged fly anatomy using X-ray microcomputed tomography (XR µCT)50,51, and segmented the volumetric image data to produce 3D reconstructions (Fig. 4A). We imaged germ-free flies and flies colonized with LpWF, Ai, or both LpWF and Ai. Numerous crypts were apparent along the length of the gut, including in uncolonized regions of the midgut and hindgut that are shielded by peritrophic matrix (Figs. 4A, S8)52. In the colonized region of the foregut, the longitudinal striations where we observed bacteria coincided with ridges and furrows of host tissue in the proventriculus inner lumen and crop duct (Fig. 4B–F). The furrows were straight in the anterior proventriculus, becoming larger and more irregular in the posterior (Fig. 4D, F). Transverse slices of the lumen wall revealed a narrow passage through the germ-free proventriculus (Fig. 4C), while the opening was much broader in the colonized proventriculus (Fig. 4E), corresponding to a significantly higher luminal volume than in germ-free flies (Fig. 4G).

A Xray µCT model of a whole fly. Cutaway shows (1) exposed proventriculus (also inset of (B)), (2) anterior midgut, and (3) posterior midgut. B Detail of proventriculus. C Cross-section of a germ-free proventriculus inner lumen. Scale bar: 5 µm. D Germ-free proventriculus inner lumen volume rendering. E LpWF-colonized proventriculus inner lumen cross-section. Scale bar: 5 µm. F LpWF proventriculus inner lumen volume rendering. G Cardia volume calculated from surface models (n = 3 to 4 flies per condition; p = 0.0025, one-way ANOVA relative to germ-free; Tukey’s correction for multiple comparisons; GF vs. Lp p = 0.020; GF vs. Lp+Ai p = 0.022.). H–M Transmission electron microscopy transverse cross-section of anterior proventriculus in (H) germ-free fly, (I) conventionally-reared fly (only lab fly bacteria; no LpWF), ( J, K) 1 hpi with LpWF, (L, M) 3 dpi colonized with LpWF (see Fig. S7). n ≥ 3 biological replicates per treatment for TEM. Yellow arrowheads indicate lumen space. Source data are provided as a Source Data file.

Consistent with XR µCT imaging, TEM cross-sections of the proventriculus of germ-free flies showed a narrow luminal space, approximately 0.5 µm in diameter (Figs 4H, S9). Similar morphology was observed in conventionally reared lab flies, which are associated with poor-colonizing strains of the same bacterial species, including L. plantarum24 (Fig. 4I). In LpWF-colonized flies, the diameter of the furrows increased to ~1 µm by 1 hpi (Figs. 4J,K, S9) and ~2–3 µm by 3 dpi (Figs. 4L, M, S9E–J), suggesting a sustained host response to niche occupancy. Heat-killed LpWF did not produce this niche expansion, indicating live LpWF cells are necessary for the luminal expansion (Fig. S7C–F). By TEM, the expanded luminal space of the colonized proventriculus contained two zones: a clear zone adjacent to the lumen wall, and a bacteria-colonized zone closer to the center of the lumen (Fig. 4L, M, S9E–J). High pressure freezing fixation showed the same phenotypes (Fig. S9S), indicating that the zonation is not simply an artifact of fixation. Taken together, our imaging results show that the proventriculus undergoes morphological changes upon colonization, which coincide with the promotion of Ai colonization.

Mucus is heavily glycosylated, with various glycan subunits including N-aceytlglucosamine, N-acetylgalatosamine, N-acetylneuraminic acid, mannose, glucose, fucose, and arabinose, depending on the type of mucus53. We hypothesized that the extracellular matrix (ECM) in the colonized zone of the proventriculus inner lumen, as seen by thin section TEM (Figs. 4, S9), is glycan-rich. To test this hypothesis, we sectioned the proventriculus and stained with a panel of lectins that have specificity for glycans found in mucus (“Methods”). Wheat germ agglutinin (WGA), Dolichus biflorus agglutinin (DBA), and Lens culinarus agglutinin (LCA) stained fly cells in the proventriculus sections, while other lectins showed no consistent binding to the proventriculus (Fig. 5A). Examining the inner lumen of the proventriculus where the secreted layer appears by TEM (Figs. 4H–M, S9), we observed staining by WGA (Fig. 5B). DBA and LCA staining was also present in the inner lumen but the locations did not correspond to the secreted layer seen by TEM (Figs. S10A, B). WGA binds N-acetylglucosamine and can also bind N-acetylneuraminic acid54, the major sialic acid found in mammalian cells. Sambucus nigra agglutinin, which binds sialic acids, did not stain, consistent with published reports that sialic acid occurs only in fly embryos55. Succinylated WGA (sWGA) binds only N-acetylglucosamine54 and exhibited a staining pattern consistent with WGA and TEM images of the secreted layer, indicating that the inner lumen ECM is N-acetylglucosamine rich (Fig. 5C). Chitin is a polymer of N-acetylglucosamine, and a permeable, chitinous cuticle is also present in the proventriculus38. Calcofluor, which binds chitin, stained the cuticle, which can be seen clearly between the inner lumen epithelial cells and the secreted layer, indicating that the secreted layer is not chitin (Fig. 5B, C). TEM suggested germ-free flies also have a narrow secreted layer (Fig. 4H). We analyzed germ-free flies and found that this narrow layer stained with WGA, indicating that N-aceytlglucosamine is a primary glycan in the secreted layer of germ-free flies (Figs. 5D, S10C). To determine whether the layer could be the product of digested chitin from yeast cell walls in the food, we sectioned newly eclosed virgin females before their first feed and found that staining of their proventriculus was consistent with germ-free flies (Figs. 5E, S10D, E). Because the proventriculus is formed from an imaginal disk during pupariation, our evidence indicates the secreted N-acetylglucosamine-rich ECM is clearly produced by the fly.

A Table of lectins that stained the proventriculus. Lectin staining of proventriculus transverse sections for B WGA in a colonized fly, C sWGA in a colonized fly, D WGA in a germ-free fly, and E WGA in a newly eclosed germ-free fly before first food ingestion. Scale bars: 20 µm. Arrowheads indicate lectin staining in the interior proventriculus lumen. n ≥ 3 biological replicates per treatment. CF: calcofluor.

Our results show that specific strains of Drosophila gut bacteria colonize crypt-like furrows in the proventriculus, that the colonization by these strains is saturable, suggesting a limited number of binding sites, and that the proventriculus responds to colonization through engorgement, which promotes colonization by bacteria that benefit the fly32,33,56. The finding that Drosophila has a specific niche for binding of commensals to sites in the crop duct and proventriculus is significant because it provides insight into how a microbiome can interact with the host in a manner that can be host-regulated and mutually beneficial. Furthermore, it predicts the existence of specific molecules in the extracellular matrix of the proventriculus that bind to the bacterial surface of colonization-competent strains but not to non-colonizing strains. Though we have not investigated mucins here, flies have mucins57,58, and three of these are expressed in the proventriculus52. Furthermore, the morphology of the extracellular matrix in the proventriculus is reminiscent of mammalian mucus, which has two layers: a dense, uncolonized layer adjacent to the epithelium, and a thinner, distal layer colonized by bacteria59. We speculate that the niche cells in the proventricular furrows either produce the adhesion substrate for the bacteria or produce an extracellular matrix that sequesters the adhesion substrate from another source, such as the salivary gland. The apparent densely stacked membranes observed by TEM of the niche cells (Fig. 1J) suggests these may be secretory cells, which is consistent with the extensive secretory nature of the proventriculus38. The finding that binding of one bacterial strain can lead to structural changes that open niche sites for a second bacterial species provides a model for how complex assemblies of bacterial strains can arise and be maintained within a host digestive tract, with the host selecting primary colonizers that in turn select secondary colonizers. Indeed, L. plantarum is homofermentative, and we have previously shown that LpWF has a positive metabolic interaction with Acetobacter strains found in the fly gut, including Ai60.

Despite the long history of studies on the Drosophila microbiome, the existence of a specific niche has been obscured by the presence of bacteria in the food and on traditional culturing media. A substantial fraction of gut bacteria under such conditions simply pass through and do not interact specifically with the gut61, even though specific microbiome members bound to their associated niches might be present. We used bacterial pulse-chase protocols to push out unbound bacteria. This methodological advance enriched for only specifically interacting cells, allowing us to identify the symbiotic niche.

Possession of a microbiome is clearly highly beneficial for Drosophila, given that axenic flies show strongly reduced growth and fecundity29,30,31,32,62,63. However, it is less clear how the relationship between the host and specific strains of bacteria is stably perpetuated. A previous study showed that larvae excrete N-acetylglucosamine in their frass, providing a nutrient that aids external L. plantarum growth on fly food37. While the origin of the larval excreta N-aceytlglucosamine was not determined, our results indicate that the adult proventriculus niche is enriched in fly-produced N-aceytlglucosamine, suggesting that the niche provides both a spatial habitat and a nutritional source for L. plantarum.

Understanding the proventricular niche is likely to provide insight into microbiome function by (1) revealing the spatial locations where bacteria influence the host to introduce molecules into the gut, perhaps along with the peritrophic membrane; and by (2) revealing whether changes in niche structure induced by one species lay the groundwork for more complex associations between different members of the microbiome, such as LpWF and Ai, that are related to their functional pathways. Finally, these observations raise the question of whether additional niches exist at other locations in the Drosophila digestive system and within the gut of other animals, including humans.

No ethical approval was obtained because insect models do not require ethical approval under local laws and regulations. All flies in this study were mated females (except in Fig. S2), which show low heterogeneity in gut morphology40. Previous work showed that the colonization phenotypes we measured are general across multiple genetic backgrounds, including CantonS, w1118, and OregonR24. Flies were reared in Wide Drosophila Vials (Cat #: 32-114, Genesee), with Droso-Plugs® (Cat #: 59-201, Genesee). Food composition was 10% glucose (filter-sterilized), 5% autoclaved live yeast, 0.42% propionic acid (filter-sterilized), 1.2% autoclaved agar, and 0.5% autoclaved cornmeal. Each vial contained 4 mL of food. Germ free fly stocks were passaged to fresh vials every 3–4 d. Five day-old mated female adults were sorted the day prior to beginning an experiment.

Liquid food was composed of 10% glucose, 5% yeast extract, and 0.42% propionic acid. The only nutritional difference between liquid and solid food was yeast extract instead of autoclaved live yeast because the yeast cell walls clog the capillaries used for liquid feeding. The bottom of capillary feeder vials contained 1.2% agar as a hydration and humidity source. Both CAFÉ- and solid food-fed flies were transferred daily to fresh vials to minimize bacterial re-ingestion. Samples of flies were surface-sterilized and crushed, and CFUs were enumerated.

Bacterial strains were reported in ref. 24, including Lactobacillus plantarum WF, L. plantarum LF, and L. plantarum WCFS1, which was called L. plantarum HS in ref. 24. Acetobacter indonesiensis SB003 was assayed for colonization in Fig. S1 of ref. 24. Fluorescent protein-expressing plasmid strains were developed and reported in refs. 24,60. pCD256-p11-mCherry, used for L. plantarum, was the generous gift of Reingard Grabherr (BOKU, Austria)64. pCM62, used for Acetobacter indonesiensis, was the generous gift of Elizabeth Skovran (SJSU, USA).

The colonization assay followed the protocol used in Fig. S1A of ref. 24. Briefly, a measured dose of bacteria was pipetted evenly on the surface of a germ-free fly food vial and allowed to absorb for 15 min. Twenty-five germ-free, 5- to 7-d post-eclosion, mated female flies were introduced to the vial and allowed to feed for a defined period of time. Flies were then removed from the inoculation vial and placed in fresh, germ-free vials. Bacteria were collected from the inoculation vial by vigorous rinsing with PBS, and the abundance was quantified by CFUs. At specified time points, CFUs in individual flies were enumerated by washing the flies 6 times in 70% ethanol, followed by rinsing in ddH2O, and then crushing and plating for CFU enumeration.

Cultures of bacteria were grown overnight in 3 mL liquid media at 30 °C. Lp strains were grown in MRS liquid media (Hardy Diagnostics, #445054), and 10 µg/mL chloramphenicol was added for mCherry-expressing strains. Ai was grown in MYPL media, and 25 µg/mL tetracycline was added for GFP-expressing strains. Bacteria were pelleted by spinning for 3 min at 400 × g, resuspended in PBS, then diluted to the desired concentration. Dose size was quantified using OD600 or by plating and counting CFUs. OD of 1.0 corresponds to 2 × 108 CFUs/mL for LpWF and 3 × 108 CFUs/mL for Ai.

Flies were inoculated by pipetting 50 µL of an appropriate concentration of the inoculum onto the food and then left to dry in the biosafety cabinet for 15 min. Flies were starved for 4 h before flipping them into the inoculation vials, where they were allowed to feed for 1 h, then flipped to fresh vials. The dose per fly was calculated as the amount of inoculum consumed divided by the number of flies in the vial. To verify that flies ate the bacteria placed on the food and measure the amount of ingested inoculum, uneaten bacteria were recovered from the vial after feeding and subtracted from the original dose. For experiments to standardize the dose of bacteria, we used an inverted 50-mL conical vial with solidified agar food in the cap, which allows for separation of food CFUs from CFUs on the walls of the vial. For other experiments, we used an autoclaved, polypropylene wide fly vial (Genesee).

Larvae were colonized by first inoculating food vials with 100 µl of bacterial culture at an OD of 1. At least 25 mated female flies were added to the vials and allowed to lay eggs for 1 day, then removed. 3 days later, 3rd instar larvae were collected and washed in PBS, then transferred to sterile agar-water vials for 4 h to clear transient bacteria. Larval guts were dissected whole, then mounted in mounting media (80% glycerol, 20% Tris 0.1 M pH 9.0), then imaged using a confocal microscope to capture a z-stack of the entire proventriculus.

Abundance in the gut was measured by homogenizing whole flies then plating to count CFUs. Flies were first anesthetized using CO2 and surface-sterilized by washing twice in 70% ethanol, then twice in PBS. Next, they were placed individually into wells of a 96-well plate along with 100 µL of PBS and ~50 µL of 0.5-µm glass beads (Biospec) and heat-sealed (Thermal Bond Heat Seal Foil, 4titude). The plate was shaken violently for 4 min at maximum speed on a bead beater (Biospec Mini-beadbeater-96, #1001) to homogenize the flies. We previously showed that the 0.5 µm bead size does not diminish bacterial counts and effectively disrupts fly tissue24. A dilution series of the entire plate was prepared using a liquid-handling robot (Benchsmart). Agar growth medium was prepared in rectangular tray plates, which were warmed and dried ~30 min prior to plating. Plates were inoculated with 2 µL of fly homogenate per well, which leads to a circular patch for CFU enumeration. The plates were incubated at 30 °C overnight. To count colonies, plates were photographed under fluorescent light and counted semi-automatically using ImageJ 2.1.065.

The number of bacteria in a fly vial was measured by recovering cells from the vial and plating on nutrient agar growth media (MRS or MYPL) to count CFUs. To collect bacteria, 2 mL of sterile PBS were pipetted into the vial. The vial was then replugged and vortexed for 10 s. A dilution series was made starting with 100 µL of the PBS wash and then plated to count CFUs. This method was used to quantify viable bacteria egested (defecated) by flies, or bacterial growth in the vial or the remainder of uneaten inoculum. Egestion and inoculation were measured over a period of 1–2 h, minimizing the opportunity for new bacterial growth.

Twelve flies were placed in a sterile polypropylene wide mouth fly vial containing 2 mL of 1.2% agar in ddH2O. Four glass capillary tubes were inserted through the flug and filled with 12 µL of filter-sterilized liquid fly food (10% glucose, 5% yeast extract, 0.42% propionic acid). Ten microliters of overlay oil were added on top to push the liquid food to the bottom of the capillary. Flies were left in the vial for 24 h before being transferred to a fresh setup. Vials were checked every 12 h to ensure flies had access to food, and a fresh flug with new capillaries was inserted if capillaries had air in them, which prevents food access. Five fly vials were put together into a 1-L beaker with a wet paper towel at the bottom and aluminum foil over the top, and the beaker was placed in the back of a fly incubator set to 25 °C, 12 h-12 h light-dark cycling, and 60% relative humidity.

LpWF bacteria were prepared by growing an overnight culture in MRS + 10 µg/mL chloramphenicol at 30 °C. The culture was pelleted by centrifugation at 400 × g and then resuspended in PBS at an OD of 2. To kill the bacteria, 1 mL of the resulting suspension in a 1.5 mL microcentrifuge tube was heated to 65 °C for 30 min in a USA Scientific Mini Dry Bath. A sample of the suspension was spread on an MRS plate and incubated for 2 days to confirm that all cells were successfully killed.

To simulate colonization by live LpWF, flies were fed with excess heat-killed bacteria daily for 3 d. 5- to 7-day old mated female flies were kept at 25 flies/vial, as in the other colonization experiments. Each day, 100 µL of the heat-killed suspension (4 × 107 CFUs/vial) was pipetted onto fresh, sterile food and allowed to dry before transferring flies into the vial. In order to clear excess killed bacteria from the niche before performing experiments to evaluate the impact on the host, flies were transferred to fresh sterile food overnight and then further cleared by placing on agar-water media for 4 h before imaging or dosing with Ai. From this point on, the methods were the same as if flies were colonized by our standard method.

To estimate the turnover time of established bacterial populations, 5- to 7-day old mated female flies were kept with 25 flies/vial. Flies were first inoculated with a pulse of fluorescently labeled, antibiotic-resistant bacteria by pipetting 50 µL of culture resuspended in PBS (OD600 = 1) onto the food and allowing it to dry prior to flipping flies into the inoculation vial. The pulse dose was allowed to establish colonization in the gut for 3 d prior to chase. Flies were fed a chase dose in the same way each day for 10 d (OD600 = 1). The abundance of labeled resident was measured daily by homogenizing and plating a sample of flies on selective media to count CFUs. The invading chase dose was assayed by plating on non-selective media. To control for any other factors that might affect resident abundance, a control group was also passaged daily to fresh food with no chase dose and assayed daily to count CFUs.

Measurements from individual flies from the different experiments were pooled by time point. Data were fit to an exponential decay using Prism, and the half-life with its confidence interval was reported.

Plasmid loss in the absence of selection was used as a proxy for bacterial growth rate24. Briefly, a standard curve was constructed by passaging plasmid-containing cells in fresh medium twice daily in a ~1:100 dilution to an OD of 0.01 for 6 d. The number of bacterial generations was estimated by counting the number of CFUs in the culture prior to dilution. The ratio of plasmid-containing CFUs to plasmid-free CFUs was counted as the number of fluorescent to non-fluorescent colonies. The doubling time is roughly 2 h for each strain. A linear regression was used to fit the standard curve data. Flies were then fed 100% plasmid-containing cells. The ratio of plasmid-containing to plasmid-free CFUs was counted at various time points, and the standard curve was used to convert the ratio to the number of doublings. In the case of dual-plasmid containing strains (Fig. S7C), growth was measured as a ratio of colonies positive for GFP-Erm plasmids (which are lost rapidly) to those positive for mCherry-Cam (which is retained much longer). A non-linear (exponential decay) regression was used. Two caveats are that (1) population bottlenecks cause wider variance in the plasmid ratio, and (2) in vivo plasmid loss rates may be different from in vitro rates. We previously showed that the first caveat can be used to infer bottlenecks. We also note that with respect to the second caveat, our use of this method to compare growth rates in a controlled experiment does not necessitate an absolute growth measurement with a standard curve. Furthermore, the growth rates in vivo were similar to in vitro, meaning that any differences in plasmid loss rates due to differences in the growth phase of the cells are likely small.

Cropectomy was performed on live flies using only new, undamaged fine forceps (#5, Dumont). Forceps, flypad, and microscope area were cleaned with 70% ethanol. Five- to 10-day old female flies were first anesthetized using CO2 then placed on a depression slide for surgery. The fly was positioned on its back, and while holding the torso with one set of forceps a small puncture was made in the abdomen just below the thorax as shown in Fig. 1O. Pressure on the forceps was released slightly to allow the tips to open up, then grab onto the crop and pull it out through the puncture. If the crop duct was still attached, it was severed along the edge of a forceps. Flies were placed in a sterile food vial and given at least 3 d to recover. Survival rate varied by operator from 1 of 10 flies to 2 of 3 flies.

Whole guts were removed from the fly by dissection with fine forceps (Dumont). Tissue was fixed in 4% PFA in PBS for 3 h at 24 °C or at 4 °C overnight. Guts were permeabilized using 0.1% Triton-X in PBS for 30 min at room temperature, washed twice in PBS, stained with 10 µg/mL DAPI for 30 min, washed twice in PBS, placed in mounting medium for up to 1 h, then transferred to the slide using a wide-bore 200 µL pipette. Each gut was then positioned on a positively charged glass microscope slide, and approximately 60 µL of mounting medium were added (mounting medium: 80% glycerol, 20% 0.1 M Tris 9.0, 0.4 g/L N-propyl gallate). Five to ten 0.1 mm glass beads (Biospec) were added to the mounting medium to form a spacer that prevents crushing of the sample. The slide was then covered with a No. 1.5 cover glass and sealed with nail polish.

Microscopy was conducted with a Leica DMi8 confocal microscope using either a 40× (1.30 NA) HC Plan Apo or a 60× (1.40 NA) HC Plan Apo oil immersion objective. Laser lines were generated using a white-light laser with AOTF crystal, and excitation wavelengths for fluorophores were: mGFP5, 488 nm; mCherry, 591 nm; Cy5, 650 nm. Whole gut images were generated by tiling multiple captures then merging using the Mosaic Merge function in the Leica Application Suite LAS X to stitch into a single stack. Z-stacks for whole guts were 70–80 µm in thickness with slices every 0.5 µm or less. To render two-dimensional images for publication, fluorescence channels were processed as maximum intensity z-projections and the brightfield channel is represented by a single z-slice from the middle of the stack.

The spatial distribution of gut colonization was quantified based on microscopy images using FIJI to mask and segment the gut regions from the images and MATLAB to quantify the extent of colonization. First, in FIJI, summed intensity z-projections of 80 µm optical sections were generated, then resized to a scale of 1 µm/px. Background subtraction with a rolling ball radius of 50 px was applied. Equivalent measurements were made on germ-free guts to quantify the autofluorescence of the gut, which varies by region.

Next, a segmented line using a spline fit and a variable width of 40–200 µm depending on the gut width of the particular region (e.g., 200 µm for the wide part of the crop and 40 µm for the proventriculus) was drawn along the length of the gut, starting with the most distal point on the crop as the origin. The “Plot Profile” function was used to measure the intensity along each of 11 segments: crop (2 segments), crop duct (2 segments), crop duct-proventriculus junction (1 segment), proventriculus (1 segment), midgut (3 segments), and hindgut (2 segments). These intensity profiles were then exported to MATLAB.

In MATLAB, the segment length for each gut region was calibrated to the average length of each segment using a bilinear fit. This step resulted in all the guts being aligned and having the same overall size so that we could compare the intensity for each region of the gut across the replicate guts. The value of each intensity value along the spline was then background subtracted and normalized to the intensity of the visually confirmed bacteria in that region. This step adjusts for the variation in autofluorescence along the gut (highest in crop and hindgut), and it adjusts for the differences in fluorescence intensity of the bacteria due to different tissue depths. After this normalization step, a 100 µm moving average filter was applied within the boundaries of each gut region to smooth the small scale spatial variation. To convert each spatial location along the gut from the intensity of colonization to colonized/uncolonized, we thresholded the intensities of each gut based on visual inspection. The proportion of all the guts with colonization at each location along the spline was then plotted in the figure panels. Additionally, the number of guts with >5% colonization within each of the delineated regions is reported as a percentage for each of the regions as follows: crop, crop duct, proventriculus, midgut, hindgut.

To measure shedding of polystyrene beads (Spherotech FP-0552-2, sky blue), flow cytometry was used to quantify the number of egested beads. Flies were kept in inverted 50 mL conical tubes with 1 mL of solid food in the cap. To collect shed material, the tubes were rinsed with 10 mL of PBS, vortexed for 10 s, and then a clean cap was placed on top. To concentrate the solution, the samples were spun in a centrifuge for 7 min at 400 × g. The pellet was then resuspended in 200 µL of PBS. The concentrated sample was counted on an Attune flow cytometer (Thermo Fisher).

Whole guts were dissected in Cacodylate pH 7.4 (Cac) buffer, then fixed for 2 d in 3% GA + 1%FA in 0.1 M Cac at 4 °C. Samples were embedded in agarose and stored at 4 °C until further processing. Samples were then washed in Cac buffer, stained with 1% OsO4 + 1.25% KfeCN for 1 h, washed in water, treated with 0.05 M maleate pH 6.5 (Mal), stained with 0.5% uranyl acetate in Mal for 1.25 h, then washed with increasing concentrations of ethanol. For embedding in resin, samples were treated with resin+propylene oxide (1:1) evaporated overnight as a transition solvent prior to embedding, then embedded in epoxy resin (Epon+Quetol (2:1)+Spurr (3:1)+2% BDMA overnight at 55 °C and cured at 70 °C for 4 d.

Samples were prepared for XR µCT following the protocol in ref. 51, which the authors generously shared prior to publication. Briefly, flies were washed in 1% Triton-X in PBS to reduce cuticular wax. A shallow hole was poked in the abdomen and thorax with a fine tungsten pin to increase permeation of fixative and stain. Fixation was with Bouin’s solution. Staining was with phoshotungstic acid for 3 weeks. Flies were mounted for imaging in a 10 µL micropipette tip containing deionized water and sealed with parafilm. Imaging was performed at the Lawrence Berkeley National Laboratory’s synchrotron Advanced Light Source on beamline 8.3.2 with assistance of Dula Parkinson. 1313 images were acquired per specimen at 20× magnification through 180 degrees of rotation. Back-projections were performed using Tomopy with the following specifications:

doFWringremoval 0 doPhaseRetrieval 1 alphaReg 0.5 doPolarRing 1 Rmaxwidth 30 Rtmax 300

Further specifications are available here: http://microct.lbl.gov/. The images in Fig. 4A, B were produced in Octopus and VG Studio 2.2. Volumetric reconstructions of the gut lumen in Fig. 4D–G were performed in Imaris using manual segmentation.

To embed flies in OCT (McKessen, 981385), the legs and wings were first removed. Next, flies were equilibrated to the OCT medium by submerging and agitating to remove bubbles. Flies were then transferred to fresh OCT in the Cryomold (Ted Pella Inc., 4565, Lot 78652). Up to 5 flies were oriented in parallel in a single mold. The block was rapidly frozen by placing it on a bed of powdered dry ice and then stored at −80 °C until sectioning. 10 µm sections were prepared at −24 °C using a Leica CM3050 cryostat and transferred onto positively charged slides (VWR Superfrost Plus, 48311-703). Sections were air dried using a Mini Dry Bath (USA Scientific, BSH200) then stored at −20 °C until staining.

Sections were fixed and stained within 2 days of sectioning. First, excess OCT was removed from the slides by briefly washing in HBSS (Cold Harbor Springs Protocol). Next, sections were fixed in 4% PFA for 30 min, then washed once in HBSS. Before staining, the sections were blocked using 1% BSA in HBSS. Sections were stained for 30 min with the specific lectin at its optimal concentration: (12.5 µg/mL Wheat germ agglutinin (WGA – Biotium, 29026-1, Lot 18W1205), 50 µg/mL Dolichos biflorus agglutinin (DBA – Glycomatrix, 21511013-1, Lot L20092804ZH), 50 µg/mL Lens culinarus agglutinin (LCA – Glycomatrix, 21511020-1, Lot L20092902ZH), or 50 µg/mL WGA-Succinylated (S-WGA – Vectorlabs, FL-1021S-5, Lot 2008145). Lectin staining was performed simultaneously with the counterstain Calcofluor (Sigma-Aldrich, 910090, Lot MKCL1227) in HBSS. Sections were then washed once in HBSS and once more in 10% HBSS. Sections were mounted in mounting media (80% glycerol, 20% Tris 0.1M pH 9.0) and covered with a #1 cover slip.

The optimal concentration for staining of each lectin was determined by preparing concentrations of 50 µg/mL, 12.5 µg/mL and 3 µg/mL in the presence of the specific haptenic sugar at a concentration of 500 mM. For WGA and S-WGA, the haptenic sugar was N-acetylglucosamine (Vectorlabs, S-9002, Lot ZJ022). For DBA, the haptenic sugar was N-acetylgalactosamine (Vectorlabs, S-9001, Lot ZJ0301). For LCA, the haptenic sugar was D-mannose (Sigma, M602025G, Lot SLBG0980V). The specificity of each lectin was assessed by the inhibition of staining by the haptenic sugar. The lowest lectin concentration that stained the tissue and could be inhibited by its corresponding haptenic sugar was chosen as the optimal concentration and used for subsequent procedures.

Statistical tests were performed in Prism. In general, data were checked for normality using a Shapiro–Wilk test. If normality was established, a Welch’s t-test was performed. Statistical tests of CFU abundances were performed on log10-transformed data. When CFUs were 0, the log was set to 0 (corresponding to a pseudocount of 1). When multiple comparisons were made, an ordinary one-way ANOVA was performed. If significant, multiple pairwise comparisons were performed with Tukey’s multiple comparisons test. When data were not normally distributed, comparisons were made using Wilcoxon rank-order tests. Error bars on proportions are either standard error of the proportion (s.e.p.), or binomial 95% confidence intervals using the Clopper–Pearson method or Jeffries method, as specified in the text. The statistical significance of differences in proportions was assessed using a Z-test.

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

All data generated or analyzed during this study are included in this published article (and its supplementary information files). Source data are provided with this paper.

Durack, J. & Lynch, S. V. The gut microbiome: relationships with disease and opportunities for therapy. J. Exp. Med. 216, 20–40 (2019).

Article CAS PubMed PubMed Central Google Scholar

Baumgartner, M. et al. Mucosal biofilms are an endoscopic feature of irritable bowel syndrome and ulcerative colitis. Gastroenterology 161, 1245–1256.e20 (2021).

Article CAS PubMed Google Scholar

Lee, J. et al. Young versus aged microbiota transplants to germ-free mice: increased short-chain fatty acids and improved cognitive performance. Gut Microbes 12, 1–14 (2020).

Article PubMed Google Scholar

Fisher, C. K. & Mehta, P. Identifying keystone species in the human gut microbiome from metagenomic timeseries using sparse linear regression. PLoS One 9, e102451 (2014).

Article ADS PubMed PubMed Central Google Scholar

Subramanian, S. et al. Persistent gut microbiota immaturity in malnourished Bangladeshi children. Nature 510, 417–421 (2014).

Article ADS CAS PubMed PubMed Central Google Scholar

Ng, K. M. et al. Microbiota-liberated host sugars facilitate post-antibiotic expansion of enteric pathogens. Nature 502, 96–99 (2013).

Article ADS CAS PubMed PubMed Central Google Scholar

Dethlefsen, L. & Relman, D. A. Incomplete recovery and individualized responses of the human distal gut microbiota to repeated antibiotic perturbation. Proc. Natl Acad. Sci. USA. 108, 4554–4561 (2011).

Article ADS CAS PubMed Google Scholar

Lawley, T. D. & Walker, A. W. Intestinal colonization resistance. Immunology 138, 1–11 (2013).

Article CAS PubMed Google Scholar

Muscariello, L., De Siena, B. & Marasco, R. Lactobacillus cell surface proteins involved in interaction with mucus and extracellular matrix components. Curr. Microbiol. 77, 3831–3841 (2020).

Article CAS PubMed PubMed Central Google Scholar

Frese, S. A. et al. Molecular characterization of host-specific biofilm formation in a vertebrate gut symbiont. PLoS Genet. 9, e1004057 (2013).

Article PubMed PubMed Central Google Scholar

Sender, R., Fuchs, S. & Milo, R. Revised estimates for the number of human and bacteria cells in the body. PLoS Biol. 14, e1002533 (2016).

Article PubMed PubMed Central Google Scholar

Faith, J. J. et al. The long-term stability of the human gut microbiota. Science 341, 1237439 (2013).

Article PubMed PubMed Central Google Scholar

Sonnenburg, E. D. et al. Diet-induced extinctions in the gut microbiota compound over generations. Nature 529, 212–215 (2016).

Article ADS CAS PubMed PubMed Central Google Scholar

Rothschild, D. et al. Environment dominates over host genetics in shaping human gut microbiota. Nature 555, 210–215 (2018).

Article ADS CAS PubMed Google Scholar

David, L. A. et al. Diet rapidly and reproducibly alters the human gut microbiome. Nature 505, 1–18 (2013).

Caporaso, J. G. et al. Moving pictures of the human microbiome. Genome Biol. 12, R50 (2011).

Article PubMed PubMed Central Google Scholar

Kim, J. K. et al. Understanding regulation of the host-mediated gut symbiont population and the symbiont-mediated host immunity in the Riptortus-Burkholderia symbiosis system. Dev. Comp. Immunol. 64, 75–81 (2016).

Article PubMed Google Scholar

Kikuchi, Y., Ohbayashi, T., Jang, S. & Mergaert, P. Burkholderia insecticola triggers midgut closure in the bean bug Riptortus pedestris to prevent secondary bacterial infections of midgut crypts. ISME J. 14, 1–12 (2020).

McFall-Ngai, M. J. The importance of microbes in animal development: lessons from the squid-vibrio symbiosis. Annu. Rev. Microbiol. 68, 177–194 (2014).

Article CAS PubMed PubMed Central Google Scholar

Fung, C. et al. High-resolution mapping reveals that microniches in the gastric glands control Helicobacter pylori colonization of the stomach. PLoS Biol. 17, 1–28 (2019).

Article Google Scholar

Lee, S. M. et al. Bacterial colonization factors control specificity and stability of the gut microbiota. Nature 501, 426–429 (2014).

Article ADS Google Scholar

Saffarian, A. et al. Crypt- and mucosa-associated core microbiotas in humans and their alteration in colon cancer patients. mBio 10, G351–20 (2019).

Article Google Scholar

Douglas, A. E. The Drosophila model for microbiome research. Lab Anim. 47, 157–164 (2018).

Article Google Scholar

Obadia, B. et al. Probabilistic invasion underlies natural gut microbiome stability. Curr. Biol. 27, 1999–2006.e8 (2017).

Article CAS PubMed PubMed Central Google Scholar

Broderick, N. A. & Lemaitre, B. Gut-associated microbes of Drosophila melanogaster. Gut Microbes 3, 307–321 (2012).

Article PubMed PubMed Central Google Scholar

Wong, C. N. A., Ng, P. & Douglas, A. E. Low-diversity bacterial community in the gut of the fruitfly Drosophila melanogaster. Environ. Microbiol. 13, 1889–1900 (2011).

Article CAS PubMed PubMed Central Google Scholar

Ludington, W. B. & Ja, W. W. Drosophila as a model for the gut microbiome. PLoS Pathog. 16, e1008398 (2020).

Article PubMed PubMed Central Google Scholar

Douglas, A. E. Simple animal models for microbiome research. Nat. Rev. Microbiol. 17, 1–12 (2019).

Walters, A. W. et al. The microbiota influences the Drosophila melanogaster life history strategy. Mol. Ecol. 29, 639–653 (2020).

Article PubMed PubMed Central Google Scholar

Lee, H.-Y., Lee, S.-H., Lee, J.-H., Lee, W.-J. & Min, K.-J. The role of commensal microbes in the lifespan of Drosophila melanogaster. Aging (Albany NY). 11, 4611–4640 (2019).

Article CAS PubMed PubMed Central Google Scholar

Téfit, M. A. & Leulier, F. Lactobacillus plantarum favors the early emergence of fit and fertile adult Drosophila upon chronic undernutrition. J. Exp. Biol. 220, 900–907 (2017).

PubMed PubMed Central Google Scholar

Consuegra, J. et al. Metabolic cooperation among commensal bacteria supports Drosophila juvenile growth under nutritional stress. iScience 23, 101232 (2020).

Article ADS CAS PubMed PubMed Central Google Scholar

Henriques, S. F. et al. Metabolic cross-feeding in imbalanced diets allows gut microbes to improve reproduction and alter host behaviour. Nat. Commun. 11, 4236 (2020).

Article ADS CAS PubMed PubMed Central Google Scholar

Gould, A. L. et al. Microbiome interactions shape host fitness. Proc. Natl Acad. Sci. USA. 115, E11951–E11960 (2018).

Article ADS CAS PubMed PubMed Central Google Scholar

Eble, H., Joswig, M., Lamberti, L. & Ludington, W. B. Cluster partitions and fitness landscapes of the Drosophila fly microbiome. J. Math. Biol. 79, 861–899 (2019).

Article MathSciNet PubMed PubMed Central MATH Google Scholar

Pais, I. S., Valente, R. S., Sporniak, M. & Teixeira, L. Drosophila melanogaster establishes a species-specific mutualistic interaction with stable gut-colonizing bacteria. PLoS Biol. 16, e2005710 (2018).

Article PubMed PubMed Central Google Scholar

Storelli, G. et al. Drosophila perpetuates nutritional mutualism by promoting the fitness of its intestinal symbiont Lactobacillus plantarum. Cell Metab. 27, 1–16 (2018).

Article Google Scholar

King, D. G. Cellular organization and peritrophic membrane formation in the cardia (proventriculus) of Drosophila melanogaster. J. Morphol. 196, 253–282 (1988).

Article CAS PubMed Google Scholar

Marianes, A. & Spradling, A. C. Physiological and stem cell compartmentalization within the Drosophila midgut. Elife 2, e00886–e00886 (2013).

Article PubMed PubMed Central Google Scholar

O’Brien, L. E., Soliman, S. S., Li, X. & Bilder, D. Altered modes of stem cell division drive adaptive intestinal growth. Cell 147, 603–614 (2011).

Article PubMed PubMed Central Google Scholar

Obniski, R., Sieber, M. & Spradling, A. C. Dietary lipids modulate notch signaling and influence adult intestinal development and metabolism in Drosophila. Dev. Cell 47, 98–111.e5 (2018).

Article CAS PubMed PubMed Central Google Scholar

Tilman, D. Competition and biodiversity in spatially structured habitats. Ecology 75, 2–16 (1994).

Article Google Scholar

Peay, K. G., Belisle, M. & Fukami, T. Phylogenetic relatedness predicts priority effects in nectar yeast communities. Proc. R. Soc. B Biol. Sci. 279, 749–758 (2011).

Amor, D. R., Ratzke, C. & Gore, J. Transient invaders can induce shifts between alternative stable states of microbial communities. Sci. Adv. 6, 1–9 (2020).

Article Google Scholar

Friedman, J., Higgins, L. M. & Gore, J. Community structure follows simple assembly rules in microbial microcosms. Nat. Publ. Gr. 1, 1–7 (2017).

Google Scholar

Martínez, I., Maldonado-Gomez, M. X. & Gomes-Neto, J. C. Experimental evaluation of the importance of colonization history in early-life gut microbiota assembly. Elife 7, e36521 (2018).

Article PubMed PubMed Central Google Scholar

Maldonado-Gómez, M. X. et al. Stable engraftment of Bifidobacterium longum AH1206 in the human gut depends on individualized features of the resident microbiome. CHOM 20, 515–526 (2016).

Google Scholar

Pitiwittayakul, N., Yukphan, P., Chaipitakchonlatarn, W., Yamada, Y. & Theeragool, G. Acetobacter thailandicus sp. nov., for a strain isolated in Thailand. Ann. Microbiol. 65, 1855–1863 (2015).

Article CAS Google Scholar

Sannino, D. R., Dobson, A. J., Edwards, K., Angert, E. R. & Buchon, N. The Drosophila melanogaster Gut Microbiota Provisions Thiamine to Its Host. mBio 9, e00155–18 (2018).

Article CAS PubMed PubMed Central Google Scholar

Mattei, A. L., Riccio, M. L., Avila, F. W. & Wolfner, M. F. Integrated 3D view of postmating responses by the Drosophila melanogaster female reproductive tract, obtained by micro-computed tomography scanning. Proc. Natl Acad. Sci. USA 112, 8475–8480 (2015).

Article ADS CAS PubMed PubMed Central Google Scholar

Schoborg, T. A., Smith, S. L., Smith, L. N., Douglas Morris, H. & Rusan, N. M. Micro-computed tomography as a platform for exploring Drosophila development. Developement 146, dev176685 (2019).

CAS Google Scholar

Buchon, N. et al. Morphological and molecular characterization of adult midgut compartmentalization in Drosophila. Cell Rep. 3, 1725–1738 (2013).

Article CAS PubMed Google Scholar

Johansson, M. E. V., Sjövall, H. & Hansson, G. C. The gastrointestinal mucus system in health and disease. Nat. Rev. Gastroenterol. Hepatol. 10, 352–361 (2013).

Article CAS PubMed PubMed Central Google Scholar

Monsigny, M., Roche, A. ‐C., Sene, C., Maget‐Dana, R. & Delmotte, F. Sugar‐lectin interactions: how does wheat‐germ agglutinin bind sialoglycoconjugates? Eur. J. Biochem. 104, 147–153 (1980).

Article CAS PubMed Google Scholar

Roth, J. et al. Occurrence of sialic acids in Drosophila melanogaster. Science 256, 673–675 (1992).

Article ADS CAS PubMed Google Scholar

Sommer, A. J. & Newell, P. D. Metabolic basis for mutualism between gut bacteria and its impact on the Drosophila melanogaster host. Appl. Environ. Microbiol. 85, e01882–18 (2019).

Article ADS CAS PubMed PubMed Central Google Scholar

Syed, Z. A., Härd, T., Uv, A. & van Dijk-Härd, I. F. A potential role for Drosophila mucins in development and physiology. PLoS One 3, e3041 (2008).

Article ADS PubMed PubMed Central Google Scholar

Huang, Y., Li, L. & Rong, Y. S. JiangShi: a widely distributed Mucin-like protein essential for Drosophila development. G3: Genes, Genomes, Genetics. 12, jkac126 (2022).

Altmann, G. G. Morphological observations on mucus-secreting nongoblet cells in the deep crypts of the rat ascending colon. Am. J. Anat. 167, 95–117 (1983).

Article CAS PubMed Google Scholar

Aranda-Díaz, A. et al. Bacterial interspecies interactions modulate pH-mediated antibiotic tolerance. Elife 9, e51493 (2020).

Article PubMed PubMed Central Google Scholar

Blum, J. E., Fischer, C. N., Miles, J. & Handelsman, J. Frequent replenishment sustains the beneficial microbiome of Drosophila melanogaster. mBio 4, e00860–13 (2013). e00860-13.

Article PubMed PubMed Central Google Scholar

Keebaugh, E. S., Yamada, R., Obadia, B., Ludington, W. B. & Ja, W. W. Microbial quantity impacts Drosophila nutrition, development, and lifespan. iScience 4, 247–259 (2018).

Article ADS CAS PubMed PubMed Central Google Scholar

Brummel, T., Ching, A., Seroude, L., Simon, A. F. & Benzer, S. Drosophila lifespan enhancement by exogenous bacteria. Proc. Natl Acad. Sci. USA 101, 12974–12979 (2004).

Article ADS CAS PubMed PubMed Central Google Scholar

Spath, K., Heinl, S., Egger, E. & Grabherr, R. Lactobacillus plantarum and Lactobacillus buchneri as expression systems: evaluation of different origins of replication for the design of suitable shuttle vectors. Mol. Biotechnol. 52, 40–48 (2011).

Article Google Scholar

Dobson, E. T. A. et al. ImageJ and CellProfiler: complements in open-source bioimage analysis. Curr. Protoc. 1, 1–19 (2021).

Article Google Scholar

Marx, C. J. & Lidstrom, M. E. Development of improved versatile broad-host-range vectors for use in methylotrophs and other Gram-negative bacteria. Microbiology 147, 2065–2075 (2001).

Article CAS PubMed Google Scholar

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Plasmid pCD256-mCherry64 was generously provided by Reingard Grabherr of BOKU, Austria. Plasmid pCM6266 was generously provided by Elizabeth Skovran of San Jose State University, USA. KCH is a Chan-Zuckerberg Biohub Investigator. Funding was provided by the Banting and the Pacific Institute for the Mathematical Sciences Postdoctoral Fellowships to EWJ, a Howard Hughes Medical Institute International Student Research fellowship, Stanford Bio-X Bowes fellowship, and the Siebel Scholars program to AA-D, the Allen Discovery Center at Stanford on Systems Modeling of Infection to KCH, the David and Lucile Packard Foundation and the Institute for Collaborative Biotechnologies through Grant W911NF-09-0001 from the US Army Research Office to JMC, a Natural Sciences and Engineering Research Council of Canada Discovery Grant and the Canada Research Chairs program to DAS, the Howard Hughes Medical Institute to ACS and CW, National Institutes of Health grants DP5OD017851 and R01DK128454 to WBL, National Science Foundation grant IOS 2032985 to WBL and KCH, National Science Foundation grant IOS 2144342 to WBL, the Carnegie Institution for Science Endowment to WBL and ACS, a Carnegie Institution of Canada grant to WBL and DAS, and National Institutes of Health training grant T32GM007231 to KA. Work by ELB and MV was supported by the Laboratory Directed Research and Development Program of Lawrence Berkeley National Laboratory under U.S. Department of Energy contract No. DE-AC02-05CH11231.

Department of Embryology, Carnegie Institution for Science, Baltimore, MD, 21218, USA

Ren Dodge, Haolong Zhu, Daniel J. Martinez, Chenhui Wang, Kevin Aumiller, Allan C. Spradling & William B. Ludington

Department of Physics, Simon Fraser University, Burnaby, BC, V5A 1S6, Canada

Eric W. Jones & David A. Sivak

Department of Physics, University of California, Santa Barbara, CA, 93106, USA

Eric W. Jones & Jean M. Carlson

Department of Biology, Johns Hopkins University, Baltimore, MD, 21218, USA

Haolong Zhu, Kevin Aumiller, Zhexian Liu, Allan C. Spradling & William B. Ludington

Molecular and Cell Biology Department, University of California, Berkeley, CA, 94720, USA

Benjamin Obadia

Howard Hughes Medical Institute, Baltimore, MD, 21218, USA

Chenhui Wang & Allan C. Spradling

Department of Bioengineering, Stanford University, Stanford, CA, 94305, USA

Andrés Aranda-Díaz & Kerwyn Casey Huang

Lawrence Berkeley National Lab, Berkeley, CA, 94720, USA

Marco Voltolini & Eoin L. Brodie

Dipartimento di Scienze della Terra, Università degli Studi di Milano, Milano, Italy

Marco Voltolini

Department of Microbiology and Immunology, Stanford University School of Medicine, Stanford, CA, 94305, USA

Kerwyn Casey Huang

Chan Zuckerberg Biohub, San Francisco, CA, 94158, USA

Kerwyn Casey Huang

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R.D., E.W.J., H.Z., B.O., C.W., E.B., J.M.C., D.A.S., A.S., and W.B.L. designed the research. R.D., E.W.J., H.Z., B.O., C.W., K.A., A.A.-D., M.V., and W.B.L. performed the research. R.D., E.W.J., D.J.M., and W.B.L. analyzed the data. R.D. and W.B.L. wrote the manuscript. R.D., E.W.J., K.C.H., D.A.S., J.M.C., A.C.S., and W.B.L. revised the manuscript. All authors reviewed the manuscript before submission.

Correspondence to William B. Ludington.

The authors declare no competing interests.

Nature Communications thanks François Leulier and the other, anonymous, reviewers for their contribution to the peer review of this work. Peer reviewer reports are available.

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Dodge, R., Jones, E.W., Zhu, H. et al. A symbiotic physical niche in Drosophila melanogaster regulates stable association of a multi-species gut microbiota. Nat Commun 14, 1557 (2023). https://doi.org/10.1038/s41467-023-36942-x

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Received: 11 January 2022

Accepted: 22 February 2023

Published: 21 March 2023

DOI: https://doi.org/10.1038/s41467-023-36942-x

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