To investigate whether communities assembled in pairs of nutrients can be predicted from those assembled in every single nutrient alone, we must first develop a quantitative null model that predicts community composition in a mixed nutrient environment in the case where each nutrient recruits species independently. Any deviation between the null model prediction and the observed (measured) composition reveals that nutrients are not acting independently, but rather ‘interact’ to shape community composition. This definition of an interaction as a deviation from a null model that assumes independent effects is commonplace in systems-level biology (Sanchez, 2019; Tekin et al., 2018).

In order to formulate the null expectation for independently acting nutrients, let us consider a simple environment consisting of two unconnected demes where two bacterial species, A and B, can grow together. The first deme contains a single growth limiting nutrient (nutrient 1), while the second deme contains a different single limiting nutrient (nutrient 2) (Figure 1A). In this scenario, each nutrient influences the abundance of species A and B independently: the microbes growing on nutrient one do not have access to nutrient two and vice versa. Let’s denote the abundance of species A in demes 1 and 2 by n_A,1 and n_A,2, and the abundance of species B as n_B,1 and n_B,2, respectively. If we now consider the two-deme environment as a whole, the abundance of species A is the sum of its abundance in each deme n_A,12 = n_A,1 + n_A,2 (likewise, for species B n_B,12 = n_B,1 + n_B,2). This example illustrates that in the scenario when two limiting nutrients act independently, each of them recruits species just as if the other nutrient were not there. In such case, the abundance of each species in a nutrient mix is the sum of what we would find in the single-nutrient habitats.

Figure 1

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Under the null model, the relative abundance of species i in a mix of nutrients 1 and 2 can be written as f_i,12(null) = w₁ f_i,1 + w₂ f_i,2 where f_i,1 and f_i,2 are the relative abundances of i in nutrients 1 and 2, respectively, and w₁ and w₂ are the relative number of cells in nutrients 1 and 2 (Materials and methods). Any quantitative difference between the null model prediction and the observed composition quantifies an ‘interaction’ between nutrients. Accounting for the presence of such interactions, the model can be re-written as f_i,12 = f_i,12(null) + ε_i,12 where ε_i,12 represents the interaction between nutrients 1 and 2 (Figure 1B).

Equipped with this null model, we can now ask to what extent the nutrients recruit species independently in mixed environments. To address this question, we followed a similar enrichment community approach to the one we have used in previous work for studying the self-assembly of replicate microbial communities in a single carbon source (Goldford et al., 2018; Estrela et al., 2020) (Materials and methods, Figure 2A). Briefly, habitats were initially inoculated from two different soil inocula. Communities were then grown in synthetic (M9) minimal media supplemented with either a single carbon source or a mixture of two carbon sources, and serially passaged to fresh medium every 48 hr for a total of 10 transfers (dilution factor = 125× ) (Figure 2A). The carbon source pairs consisted of a focal carbon source mixed at equal C-molar concentrations with one of eight additional carbon sources. We previously found that stable multi-species communities routinely assemble in a single carbon source (which is limiting under our conditions), and they converge at the family level in a manner that is largely governed by the carbon source supplied, while the genus or lower level composition is highly variable (Goldford et al., 2018). We chose glucose as the focal carbon source because we have previously carried out multiple assembly experiments in this nutrient (Goldford et al., 2018; Estrela et al., 2020). As the additional carbon sources, we chose nutrients that are simple and metabolically diverse (sugar vs acid, that contain a different number of atoms of carbon, and that enter metabolism at different points), namely ribose, fructose, cellobiose, and glycerol (i.e. a pentose, a hexose, a disaccharide, and a sugar alcohol) and fumarate, benzoate, glutamine, and glycine (two organic acids and two amino acids). All carbon sources were also used in single carbon source cultures.

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Communities assembled in single sugars contained 5–24 exact sequence variants (ESVs), mainly belonging to the Enterobacteriaceae family (mean relative abundance ± SD of~0.98 ± 0.03), a sugar specialist (Figure 2—figure supplement 1). In contrast, communities assembled in organic acids exhibited a higher richness (12–36 ESVs), and unlike in sugars, Enterobacteriaceae were generally rare (mean ± SD ~ 0.06 ± 0.06). Instead, communities were dominated by respirative bacteria mainly belonging to the Pseudomonadaceae (mean ± SD ~ 0.51 ± 0.25), Moraxellaceae (mean ± SD ~ 0.18 ± 0.21), and Rhizobiaceae (mean ± SD ~ 0.11 ± 0.13) families (Figure 2—figure supplement 1). Because of the observed family-level convergence across carbon sources, which is consistent with previous studies (Goldford et al., 2018; Estrela et al., 2020; Lu et al., 2018), we focus our analysis below on family-level abundance.

To investigate the predictive power of the null (additive) model, we compare the predicted and observed relative abundances of each family for each carbon source pair across all experiments. Our results show that the null model predicts reasonably well the family-level abundances on average (Pearson’s R = 0.95 and p<0.001; RMSE = 0.073, N = 223) (Figure 2B, Figure 2—figure supplements 2 and 3). To confirm that the strong predictive power of the null model is not an idiosyncrasy of using glucose as the focal carbon source in the pairs, we repeated the same experiment with succinate (an organic acid) as the focal carbon source. Although the correlation between observed and predicted abundance is lower than for glucose, the null additive model is still predictive (Pearson’s R = 0.87 and p<0.001; RMSE = 0.094; N = 257) (Figure 2B).

This result seems to indicate that, at the family level, a simple model that assumes that nutrients act independently can predict community composition in a pair of nutrients (for an analysis of this point at the genus and ESV level, see Figure 2—figure supplement 4). However, when we looked at this more closely and broke down our results by carbon source and family, we found consistent and systematic deviations from the null model (Figure 2C). For example, across all succinate–sugar pairs, Enterobacteriaceae are significantly more abundant than predicted by the null model (one-tailed paired t-test, N = 8, p<0.05 based on 1000 permutations; see Materials and methods), while both Rhizobiaceae and Moraxellaceae are less abundant than predicted (one-tailed paired t-test, N = 8, p<0.05 based on 1000 permutations; see Materials and methods) (Figure 2C). The null ‘interaction-free’ model also predicts species abundance better in certain carbon source combinations (e.g. glucose + ribose) than in others (e.g. glucose + glutamine) (Figure 2C). The existence of systematic deviations from the null prediction reveals that some nutrient pairs do not recruit families independently, but instead ‘interact’ with each other to affect the abundance of specific families.

To map the regularities in nutrient interactions observed, we next sought to characterize the nature of these interactions for each carbon source pair and every family. One helpful way of visualizing nutrient interactions is to draw the pairwise abundance landscape for each species and carbon source pair (Figure 3A). For instance, a species could be either more abundant in a pair of nutrients than it is in any of them independently (synergy). Or it could be less abundant than it is in any of the two (antagonism). Dominance is a less extreme interaction that can be visualized by the pushing of a species abundance toward the value observed in one of the two nutrients and away from the average, thus overriding the effect of the second available nutrient (Figure 3A).

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When the interaction is positive (ε > 0), the dominant nutrient is the one where the family grew to a higher abundance. When the interaction is negative (ε < 0), the dominant nutrient is the one where the species grew less well. Mathematically, dominance occurs when |ε| > 0 and min(f_i,1, f_i,2) ≤ f_i,12 ≤ max(f_i,1, f_i,2), while synergy and antagonism (forms of super-dominance) occur when |ε| > 0 and f_i,12 > max(f_i,1, f_i,2) and f_i,12 < min(f_i,1, f_i,2), respectively (Materials and methods). Figure 3B shows representative examples of dominant carbon source interactions. For instance, Moraxellaceae and Rhizobiaceae grow strongly on succinate, but they are not found in fructose. When fructose is mixed with succinate, both families drop dramatically in abundance, despite their high fitness in succinate alone. Interestingly, however, the dominance of fructose over succinate is not observed for all families: those two nutrients do not interact on Pseudomonadaceae, whose abundance is well predicted by the null model. Using this framework, we then systematically quantified the prevalence of dominance, antagonism, and synergy between nutrients for each family (Figure 3—figure supplement 1A). While 66% of the nutrient pair combinations exhibited no significant interaction, dominance was by far the most common interaction amongst those that interacted (75%, Figure 3—figure supplement 1A). It occurred predominantly in the sugar–acid pairs, and to a lesser extent in the acid–acid pairs, and only rarely in the sugar–sugar pairs (Figure 3—figure supplement 1B). This result strongly suggests that nutrient interactions are not random but do have a specific structure that is conserved at the family level (Figure 3—figure supplement 1C).

To systematically characterize and quantify nutrient dominance, we developed a dominance index (δ) (Materials and methods). For visualization purposes, the dominance index for the sugar–acid pairs (we will discuss the acid–acid pairs later) is written as δ_i=−|ε₁₂| when the sugar dominates and as δ_i=|ε₁₂| when the acid dominates. If ε₁₂ = 0, then δ_i=0. That is, in the absence of interaction between nutrients, there is no dominance. By plotting the dominance index for each pair of nutrients and each family, we observe a generic pattern of dominance of sugars over acids (Figure 3C). The families Moraxellaceae or Rhizobiaceae are recruited to the community by most organic acids in isolation, but they are not found in most sugar communities. When sugars and organic acids are mixed together, the sugar dominates and both families are at much lower abundances (by ~6-fold in the case of Moraxellaceae and ~114-fold in Rhizobiaceae) than expected by the null model, even though the organic acid where they thrived is present in the environment. Consistent with this pattern, we found that pairs of more-similar nutrients (a pair of sugars or a pair of organic acids) were significantly better predicted by the null model than mixed organic acid–sugar pairs (Figure 3D). No generic pattern of dominance was observed in the acid–acid mixtures (Figure 3—figure supplement 2). When we examine interactions and dominance at the genus level, we find that sugars do not exhibit the same dominance for all genera within the same family (Figure 3—figure supplements 3–4). This result is consistent with the convergence of community structure at the family level (despite substantial variation at lower levels of taxonomy), which we have reported for communities assembled in a single nutrient (Goldford et al., 2018; Estrela et al., 2020). Together, these results indicate that interactions between nutrients are not universal, but rather they are conserved at the family-level.

Our findings pose intriguing questions about the mechanisms behind the nutrient interaction patterns we have observed. For instance, is it reasonable to expect that the additive null model should have worked as well as it did, and better at the family than at the species level? Why are pairs of more-similar nutrients better explained by the null model than pairs of more dissimilar nutrients? What may explain why nutrients dominate over others at the family level? And why do sugars generally dominate organic acids for most families?

We have previously shown that many of the properties of our experimental enrichment communities reflect the generic emergent behavior of consumer-resource models (Goldford et al., 2018; Marsland et al., 2019), and subsequent work extended this finding to complex natural communities (Marsland et al., 2020a). We thus sought to ask whether our observations regarding the assembly of communities in pairs of resources are similarly reflecting a generic emergent property of consumer-resource models. To address this question, we followed the same procedure as we and others have done in previous work (Goldford et al., 2018; Marsland et al., 2020a; Marsland et al., 2020b; Serván and Allesina, 2020), and simulated the top-down assembly of microbial communities in pairs vs single nutrients using a recently developed Microbial Consumer Resource Model (MiCRM) (Goldford et al., 2018; Marsland et al., 2019; Marsland et al., 2020a) (see Materials and methods). The MiCRM differs from the classical MacArthur-Levins model (MacArthur, 1970) in that it includes metabolic cross-feeding in a manner that preserves thermodynamic balance. The model and the details of the simulations are described in the Materials and methods section. In brief, 200 species are seeded into each habitat at the start of a simulation. Each of these is represented by a different vector of resource uptake rates. These vectors are randomly sampled in a manner that captures the existence of two functional guilds, each of which specializes in a different group of resources (e.g. sugars vs organic acids) (Figure 4A). Members of the family specializing on sugars (i.e. the Enterobacteriaceae) have on average a higher uptake rate on each sugar whereas members of the family specializing on acids (i.e. the Pseudomonadaceae) have on average a higher uptake rate on each acid. The magnitude of specialization by each family on its preferred resource type is tuned by two parameters, q_A and q_S, which modulate the mean and variance of the probability distribution from which the uptake rates are sampled (see Materials and methods for more details). We note that this specialization is quantitative rather than discrete, as all species are assumed to be able to consume all of the resources (a point that is in general consistent with our experimental findings [Figure 4—figure supplement 1]). Communities are allowed to find a dynamical equilibrium, at which point we stop the simulation. In total, and in order to get to generic behavior, we generated 100 simulations each with a different random set of species (Materials and methods).

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A generic property of these simulations is that the species-level community composition on mixtures of two limiting nutrients is reasonably well described by the additive null model (Pearson’s R = 0.7 and p<0.001; RMSE = 0.097; N = 2440) (Figure 4B; Materials and methods), which is consistent with previous consumer-resource modelling work (Marsland et al., 2020a). In addition, when we group species by the functional groups they belong to (i.e. family), the predictive ability of the additive null model improves (Pearson’s R = 0.99 and p<0.001; RMSE = 0.03; N = 414) (Figure 4B), a point that is consistent with our experimental findings (Figure 2—figure supplement 4). This family-level additivity holds when communities are randomly colonized by a different set of species (Figure 4—figure supplement 2), suggesting that family-level additivity is robust to species-level taxonomic variability. The predictive accuracy of our null model is, however, influenced by the level of resource specialization. The less specialized (i.e. more generalist) the families are, the lower the predictive power of the null additive model (Figure 4—figure supplement 3).

By contrast, the simulated communities do not exhibit any systematic dominance, neither at the species nor at the family level (Figure 4B). What feature of the MiCRM might be causing us to miss this experimental behavior? One assumption of the model, which we had made for the sake of simplicity and for consistency with previous work, is that all nutrients are equally valuable for the microbes that specialize on them. In other words, the benefits of growing in each type of nutrient are symmetric. Yet, this assumption is not really consistent with the empirical reality that glucose specialists, such as Enterobacteriaceae, grow more strongly in sugars than organic acid specialists do on organic acids. This is illustrated in Figure 4C, where we plot the growth advantage for seven Enterobacteriaceae isolates in sugar media vs the growth advantage of Pseudomonadaceae, Rhizobiaceae, and Moraxellaceae isolates in organic acids.

We postulated that including this asymmetry may unbalance the competition for resources and give rise to nutrient dominance at the family level, as the family that lies on the winning side of that asymmetry may leverage its enhanced competitive ability in the most valuable nutrient to displace the losing family from its lower-value nutrient niche. To test this intuition, we relaxed the symmetry in resource value that was imposed by default in the model, and repeated our simulations for different levels of nutrient value asymmetry (the simulations still include facilitation via metabolite secretion, as we had done in all prior simulations) (Figure 4A, Figure 4—figure supplement 4). As we show in Figure 4D, and consistent with our intuition, nutrient dominance at the family level may emerge as a generic property of microbial consumer-resource models when a nutrient is substantially more valuable than the other. Reassuringly, our experiments indicate that dominance is generally favorable to the taxa that benefits from growth asymmetry for example to Enterobacteriaceae in sugar–acid mixes, and unfavorable to families in the losing end of growth asymmetry (Pseudomonadaceae, Rhizobiaceae, and Moraxellaceae) in those same environments. This observation is consistent with the behavior of the model (Figure 4D).

Our consumer-resource model shows that dominance is a general outcome of consumer-resource interactions when there exists an asymmetry in nutrient benefit (an asymmetry that is indeed observed for the families in our communities), but other mechanisms of dominance may be at play too. For instance, one plausible mechanism that could lead to dominance is oxygen limitation, in particular if different carbon sources were to have different oxygen requirements (Hempfling and Mainzer, 1975; Skrinde and Bhagat, 1982). To explore this idea of asymmetric oxygen demands, we used flux-balance analysis (FBA) to determine the oxygen demands of growth on each of the single-carbon sources. We found that, except for benzoate, all carbon sources have similar oxygen demands (Figure 4—figure supplement 5). This does not rule out, however, the possibility that kinetics of growth and oxygen uptake may still contribute to oxygen depletion in a manner that may further stimulate dominance (in addition to the asymmetric resource benefits we report in Figure 4).

Let us consider a simple scenario of two co-cultures of species A and B growing together in two separate demes, each containing a single nutrient (labeled 1 and 2). The fractions of A and B in nutrient/deme one are f_A,1 = n_A,1 /(n_A,1 + n_B,1) and f_B,1 = n_B,1/(n_A,1 + n_B,1), respectively, and similarly, the fractions of A and B in nutrient/deme two are f_A,2 = n_A,2/(n_A,2 + n_B,2) and f_B,2 = n_B,2 /(n_A,2 + n_B,2) (where n is the total number of cells of species A or B). If we consider the two-deme system as a whole (i.e. if we pool together the amount of species in each nutrient/deme), the fractions of A and B in the mixture are given by: f_A,12 = (n_A,1 + n_A,2)/(n_A,1 + n_B,1 + n_A,2 + n_B,2) and f_B,12 = (n_B,1 + n_B,2)/(n_A,1 + n_B,1 + n_A,2 + n_B,2).

We can define n_t,1 = n_A,1 + n_B,1 and n_t,2 = n_A,2 + n_B,2 as the total number of cells in the nutrient demes 1 and 2, respectively. We can thus write f_A,12 = (n_A,1 + n_A,2)/(n_t,1 + n_t,2). Defining w₁ = n_t,1/(n_t,1 + n_t,2) and w₂ = n_t,2/(n_t,1 + n_t,2), it is straightforward to show that: f_A,12 = w₁ f_A,1 + w₂ f_A,2. By the same reasoning, we find that f_B,12 = w₁ f_B,1 + w₂ f_B,2.

Soil samples were collected from two different natural sites in West Haven (CT, USA), with sterilized equipment, and placed into sterile bottles. Once in the lab, 5 g of each soil sample were then transferred to 250 mL flasks and soaked into 50 mL of sterile 1× phosphate buffer saline supplemented with 200 μg/mL cycloheximide (Sigma, C7698) to inhibit eukaryotic growth. The soil suspension was well mixed and allowed to sit for 48 hr at room temperature. After 48 hr, samples of the supernatant solution containing the ‘source’ soil microbiome were used as inocula for the experiment or stored at −80°C with 40% glycerol.

Carbon source (CS) stock solutions (Supplementary file 1a) were prepared at 0.7 C-mol/L (10×) and sterilized through 0.22 µm filters (Millipore). Carbon sources were aliquoted into 96 deep-well plates (VWR) as single CS or mixed in pairs at 1:1 (vol:vol) and stored at −20°C. The carbon sources were adjusted to equal C-molar concentrations because carbon is the main limiting factor. To keep the total amount of carbon constant across all treatments, pairs contained half the amount of each carbon source compared to their respective single CS. Synthetic minimal growth media was prepared from concentrated stocks of M9 salts, MgSO₄, CaCl₂, and 0.07 C-mol/L (final concentration) of single or pairs of CS. The final pH of all growth media is shown in Supplementary file 1a.

Starting inocula were obtained directly from the ‘source’ soil microbiome solution by inoculating 40 μL into 500 μL culture media prepared as indicated above. For each sample and carbon source, 4 μL of the culture medium was dispensed into fresh media plates containing the different single or pairs of CS in quadruplicate. Bacterial cultures were allowed to grow for 48 hr at 30°C in static broth in 96 deep-well plates (VWR). After 48 hr, each culture was homogenized by pipetting up and down 10 times before transferring 4 μL into 500 μL of fresh media, and cells were allowed to grow again. Cultures were passaged 10 times (~70 generations). OD620 was measured after 48 hr growth. Samples were frozen at −80°C with 40% glycerol.

Samples were centrifuged for 40 min at 3500 rpm, and the pellet was stored at −80°C until DNA extraction. DNA extraction was performed with the DNeasy 96 Blood and Tissue kit for animal tissues (QIAGEN), as described in the kit protocol, including the pre-treatment step for Gram-positive bacteria. DNA concentration was quantified using the Quan-iTPicoGreen dsDNA Assay kit (Molecular Probes, Inc), and the samples were normalized to 5 ng/µL before sequencing. The 16S rRNA gene amplicon library preparation and sequencing were performed by Microbiome Insights, Vancouver, Canada (https://microbiomeinsights.com/). For the library preparation, PCR was done with dual-barcoded primers (Kozich et al., 2013), targeting the 16S V4 region, and the PCR were cleaned up and normalized using the high-throughput SequalPrep 96-well Plate Kit. Samples were sequenced on the Illumina MiSeq using the 300 bp paired-end kit v3.chemistry.

The taxonomy assignment was performed as described in previous work (Estrela et al., 2020). Following sequencing, the raw sequencing reads were processed, including demultiplexing and removing the barcodes, indexes, and primers, using QIIME (version1.9, [Caporaso et al., 2010]), generating fastq files with the forward and reverse reads. DADA2 (version 1.6.0) was then used to infer ESVs (Callahan et al., 2016)​. Briefly, the forward and reverse reads were trimmed at position 240 and 160, respectively, and then merged with a minimum overlap of 100 bp. All other parameters were set to the DADA2 default values. Chimeras were removed using the ‘consensus’ method in DADA2. The taxonomy of each ESV was then assigned using the naïve Bayesian classifier method​ (Wang et al., 2007) and the Silva reference database version (Quast et al., 2013) as described in DADA2. The analysis was performed on samples rarefied to 10,779 reads.

We used OD620 after the 48 hr growth cycle as a proxy for total population size (community biomass) (Figure 2—figure supplement 5). The predicted relative abundance of species i in a mix of nutrients 1 and 2 was then calculated as f_i,12(null) = w₁ f_i,1 + w₂ f_i,2 where f_i,1 and f_i,2 are the relative abundances of i in nutrients 1 and 2, respectively, and w₁ = (OD620₁/(OD620₁ +OD620₂)) and w₂ = (OD620₂/(OD620₁ +OD620₂)). In Figures 2 and 3D, Figure 2—figure supplements 3–4, f_i,12(null) is calculated as the mean of the two single carbon source-replicate pairwise combinations (N = 16). Pearson’s R was calculated using the R function ‘cor.test’ from the ‘stats’ package, and the RMSE was calculated using the ‘rmse’ function from the ‘Metrics’ package.

To determine whether an interaction between nutrients exists (i.e. ε ≠ f_i,12 - f_i,12(null)), we assess whether the abundance observed in the carbon source mixture is significantly greater or lower than the abundance predicted by the null additive model (i.e. ε > 0 or ε < 0, respectively) (one-tailed paired t-test). More specifically, considering the two inocula and four replicates per carbon source, the family-level analysis was done as follow. For each carbon source pair and inoculum, four predicted pairs are formed by randomly pairing one replicate of each carbon source. Four unique observed vs predicted pairs are then randomly formed from the 64 possible combinations (i.e. from the four single nutrient 1 × four single nutrient 2 × four mixed nutrient 12). Up to this point, the pairs are formed for each inoculum separately, in other words, there is no cross-inocula pairing. Once all N=8 pairs are formed (i.e. N = 4 pairs per inoculum), they are pooled to perform the one-tailed paired t-test. The N = 8 pairs are then randomly permuted 1000 times, determining the t-statistic for each permutation. We establish a 95% confidence threshold for the t-statistic. The effect observed is statistically significant (i.e. an interaction exists) if a significant difference is found in more than 95% of the permuted pairs. At the genus level, the analysis was performed in a similar way, except that the two inocula were kept separately. This is because, compared to families, the likelihood that genera that are sampled in one of the inocula are sampled in the other inoculum is much lower.

Once an interaction has been identified (i.e. |ε| > 0), we can determine the type of interaction formed (Figure 3A). Synergy and antagonism (which are forms of super-dominance) occur when f_i,12 > max(f_i,1, f_i,2) and f_i,12 < min(f_i,1, f_i,2), respectively, while dominance occurs when min(f_i,1, f_i,2) <= f_i,12 <= max(f_i,1, f_i,2) (Welch two sample t-test, p<0.05). When ε > 0, the nutrient with greater abundance dominates; when ε < 0, the nutrient with lower abundance dominates. For visualization purposes, we developed a dominance index (δ). The dominance index for the sugar–acid pairs is written as δ_i = −|ε₁₂| when the sugar dominates and as δ_i = |ε₁₂| when the acid dominates. The dominance index for the sugar–sugar and acid–acid pairs is written as δ_i = −|ε₁₂| when the focal carbon source (glucose or succinate) dominates and as δ_i = |ε₁₂| when the additional carbon source dominates.

Several communities (transfer 10) from different inocula and carbon sources were plated on chromogenic agar (HiCrome Universal differential Medium, Sigma) and grown for 48 hr at 30°C. Single colonies exhibiting distinct morphologies and/or colours were picked, streaked a second time on fresh chromogenic agar plates for purity, and grown for 48 hr at 30°C. A single colony was then picked from each plate and grown into Tryptic Soy Broth (TSB) for 48 hr at 30°C. The single-strain cultures were stored with 40% glycerol at −80°C. The isolated strains were sent for full-length 16S rRNA Sanger sequencing (Genewiz), and their taxonomy was assigned using the online RDP naïve Bayesian rRNA classifier version 2.11.

Twenty-two isolated strains belonging to the four dominant families, namely Enterobacteriaceae (7), Pseudomonadaceae (5), Moraxellaceae (6), and Rhizobiaceae (4) (Supplementary file 1b), were streaked from frozen stock on chromogenic agar plates and grown for 48 hr at 30°C. For each strain, a single colony was pre-cultured in 500 µL TSB in a deepwell plate for 24 hr at 30°C. Each strain was then acclimated into the 10 single carbon sources (glucose, fructose, cellobiose, ribose, glycerol, succinate, fumarate, benzoate, glutamine, and glycine). For this, 2 µL of the grown pre-culture was inoculated into 500 µL of fresh minimal media with each carbon source at a concentration of 0.07 C-mol/l and grown for 48 hr at 30°C. The growth curve assay was then performed in a 384-well plate by inoculating 1 µL of the grown isolate culture on 100 µL of fresh media of the same carbon source as for the acclimation step (three to four replicates each). OD620 was read every 10 min for ~40 hr at 30°C. The average growth rate of each strain in each carbon source was calculated as r_avg = log₂(N_f/N_i)/(t_f−t_i) where N_f is the OD at 16 hr (ie. t_f) and N_i is the OD at 0.5 hr (i.e. t_i).

The growth rate asymmetry on sugars (q_S) is calculated as q_S = r_avg(E, S) − r_avg(O, S) where r_avg(E, S) is the mean average growth rate of Enterobacteriaceae on the sugar S, and r_avg(O, S) is the mean average growth rate of one of the other dominant families (i.e. Pseudomonadaceae, Moraxellaceae, or Rhizobiaceae) on S. The growth rate asymmetry on organic acids (q_A) is calculated as q_A = r_avg(O, A) − r_avg(E, A) where r_avg(O, A) is the mean average growth rate of one of the other dominant families (i.e. Pseudomonadaceae, Moraxellaceae, or Rhizobiaceae) on the organic acid A, and r_avg(E, A) is the mean average growth rate of Enterobacteriaceae on A.

Microbial community assembly is modeled using the Microbial Consumer Resource Model (MiCRM), with simulations implemented using Community Simulator, a freely available Python package (Marsland et al., 2020b). This model has been outlined extensively in previous work and has been shown to qualitatively reproduce ecological patterns across both natural (Goldford et al., 2018) and laboratory (Marsland et al., 2020a) microbiomes. Here we describe the exact equations simulated and parameters used in this paper. A more general description of this model is given elsewhere (Marsland et al., 2019; Marsland et al., 2020b). Our MiCRM simulations model the abundance N_i of n species and the abundance R_α of M resources in a well-mixed chemostat-like ecosystem with continuous resource flow. We focus on continuous resource flow for simplicity and because previous work has shown that the major qualitative features of the MiCRM are unaffected by periodic resource supply (as was the case in our experiments) (Marsland et al., 2020a). Species interact by uptake and release of resources into their environment. The dynamics of the system are governed by the following set of ordinary differential equations:

Here c_iα is the uptake rate of resource α by species i, m is the minimal energy requirement for maintenance of species i, τ is the timescale for supply of external resources, $R_{\alpha }^0$ is the abundance of resource α supplied (i.e. the abundance in the media), l is the fraction of resource secreted as by-product, and D_α,_β is the fraction of resource α secreted as by-product β. In line with previous work, the following parameters are kept constant for all simulations τ = 1, m = 1, and l = 0.5 (Goldford et al., 2018; Marsland et al., 2020a).

In the MiCRM, by-product production is encoded in the metabolic matrix D where each element D_α,_β specifies the fraction of resource α secreted as by-product β. As in previous work, each column β in D_α,_β is sampled from a Dirichlet distribution with concentration parameter d_α,_β=1/(sM) where s = 0.3 is a parameter that tunes the sparsity of the underlying metabolic network. The Dirichlet distribution ensures that each column sums to one so that the total secretion flux does not exceed the input flux. For simplicity we used a fixed concentration parameter and so are not assuming any underlying metabolic structure. The MiCRM also assumes that all species have the same D matrix, that is when growing on the same resource each species releases the same metabolic by-products.

In our simulations, species differ solely in the uptake rate for different resources c_iα where i is the species and α is the resource. Taxonomic specialization is introduced in the form of two families F_A and F_S that each have a preference for one of two resource classes A and S, respectively. Each c_iα is sampled from a gamma distribution (to ensure positivity) whose mean <c_iα> and variance var(c_iα) depends on the family of i and the resource class of α. This means that all species are capable of metabolizing all resources. Specifically

and

μ_c = 10 determines the overall mean uptake rate and ${\displaystyle {\sigma }_c^2}$ = 3 determines overall variance in uptake rate (these parameters are the default value in the Community Simulator package). Parameters q_A and q_S tune the relative advantage each specialist family has on its preferred resource. When q_A = 1, only F_A consumes resources in A whereas when q_A = 0 both families have equal access to resources in A. Conversely, when q_S = 1, only F_S consumes resources in S whereas when q_S = 0 both families have equal access to resources in S.

For each simulation we consider 200 species (100 per family). Each community in one simulation is seeded with all 200 species. This means that there is no stochasticity in colonization (though see Figure 4—figure supplement 2 where this assumption is relaxed). We choose 200 species as this is within the range of the number of ESVs in a typical inoculum for our experiments (110–1290 ESVs, reported in Goldford et al., 2018). The initial abundances are all set to 1 for simplicity. In line with our experiments, either one or two resources are supplied in the media and the rest are generated as metabolic by-products. For simulations with a single supplied resource, $R_{\alpha }^0$ = 1000 if α is the supplied resource and 0 otherwise. For simulations with two supplied resources, $R_{\alpha }^0$ = 500 for each supplied resource and 0 otherwise. This ensures that the total amount of resources is kept constant as in our experiments. In total, we consider 20 resources in each simulation (with 10 resources in each resource class [A or S]) as this gives us communities with 7 ± 2 species (mean ± SD) at equilibrium, which is comparable to the diversity of our experimental communities.

In line with our experiments, each simulation consisted of three types of mixed-resource environments (one with two supplied resources in class R_A, one with two supplied resources in class R_S and one with one resource in class R_A and one resource in class R_S). We also included all four single resource environments needed to predict the mixes (i.e. two with the resources in class R_A and two with the resources in class R_S). Therefore, each simulation consisted of seven communities each in a different environment and all seeded with the same initial set of 200 species. The equilibrium for all seven communities was found using the SteadyState function in Community Simulator (Marsland et al., 2020b). Failed runs where the SteadyState function returned an error were removed from our analysis. In addition, for each simulation we tested that the SteadyState algorithm had truly converged to an equilibrium using the same approach as in Marsland et al., 2020a and removed all non-convergent runs (defined as a run for which ${\displaystyle \left|dln(N_i/dt)\right|>{10}^{-5}}$). Including these runs would not have qualitatively changed our results.

In the raw numerical output of the run, all species have non-zero abundances due to limits in numerical precision. A species was considered extinct if its abundance was less than 10⁻⁶, which was set by looking at a histogram of the raw output of our simulations. Once the extinct species were removed, we predicted the relative abundance of each species i in the mixture of nutrients using the same approach that had been used for the experimental data. To obtain a statistically robust sample size, we repeated this procedure for 100 replicate simulations, resampling all randomly generated parameters in each simulation (i.e. resampling all c_iα and D_α,_β as described above).

We use FBA to estimate whether the different carbon sources were likely to result in large differences in oxygen demand. FBA is a widely used constraint-based modelling approach that allows us to predict metabolic fluxes through a stoichiometric metabolic network (assuming optimal growth and that cells are in a steady state) (Orth et al., 2010). For this analysis, we used a modified version of iJO1366 (see below), a high-quality genome scale-metabolic network of E. coli (Orth et al., 2011). FBA simulations were performed using the COBRApy Package (Ebrahim et al., 2013). We simulated the growth of E. coli on minimal synthetic media in aerobic conditions containing one of the 10 carbon sources used in our experiments (Figure 2A). These simulations were used to estimate the number of O₂ molecules that would be consumed per carbon atom when growing on each of the 10 carbon sources (Figure 4—figure supplement 5). Except for benzoate, we found that all of the carbon sources exhibited similar predicted oxygen demands (0.25–0.34 O₂/C). This does not rule out the possibility that the kinetics of growth and O₂ uptake may contribute to increased O₂ depletion in one carbon source compared to another, nor that the different taxa selected for by the different carbon sources might display differences in O₂ uptake. Nonetheless, they do suggest that differences in oxygen stoichiometry are unlikely to be the main mechanism for dominance across all carbon sources.

For these simulations, all inorganic compounds were assumed to be in excess and their exchange fluxes were unbounded by setting to an arbitrarily large negative value (−1000 mmol/gDWh). These compounds are as follows: ca2_e, cbl1_e, cl_e, co2_e, cobalt2_e, cu2_e, fe2_e, fe3_e, h_e, h2o_e, k_e, mg2_e, mn2_e, mobd_e, na1_e, nh4_e, ni2_e, pi_e, sel_e, slnt_e, so4_e, tungs_e, zn2_e, and o2_e. To estimate the optimal oxygen consumption per mole of carbon consumed, the exchange flux for each of the 10 carbon sources was set to −1 cmol/gDWh. We set the lower bound on ATPM maintenance to 0 as we wanted to estimate the O₂/C when resources were in excess and so the effects of growth independent maintenance would be negligible. Similar results can be obtained using the default ATPM lower bound in the published model and setting a higher lower bound on the carbon uptake flux (such as the −60 cmol/gDWh typically used for E. coli on glucose) (Harcombe et al., 2014). The biomass reaction (BIOMASS_Ec_iJO1366_core_53p95M) was used as the objective function.

In the absence of any additional reactions, FBA predicts that E. coli can grow on all carbon sources aside from benzoate and cellobiose. To obtain growth on cellobiose we added a cellobiose PTS transporter (CELBpts) and a beta-glucosidase (BGLA1) as well the corresponding exchange and periplasm diffusion reactions (EX_cellb_e and CLBtex). While most wild-type E. coli are unable to use cellobiose, other Enterobacteriaceae such as Klebsiella oxytoca can (using the reactions we have added) (Sekar et al., 2012). To obtain growth on benzoate, we added the benzoate degradation pathway (the β-ketoadipate pathway) from Pseudomonas putida. This included the following reactions (EX_bz_e, BZt, BZ12DOX, CATDOX, MUCCYCI, MUCLI, OXOAEL, 3OADPCOAT) (Sudarsan et al., 2016). We note that the lack of growth by E. coli on benzoate is consistent with the low abundance of other Enterobacteriaceae in the benzoate communities and the prevalence of Pseudomonas (Figure 2—figure supplement 1). The O₂/C ratio obtained on benzoate using the E. coli model (0.560) is similar to the ratio obtained using the P. putida iJN1463 model (0.548) in which the benzoate degradation pathway is native (Nogales et al., 2020).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We want to thank members of the Sanchez lab for helpful discussions. This work was supported by the National Institutes of Health through grant 1R35 GM133467-01, and by a Packard Foundation Fellowship to AS.

1. Received: December 20, 2020
2. Accepted: April 2, 2021
3. Version of Record published: April 20, 2021 (version 1)

© 2021, Estrela et al.

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