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NEWS AND VIEWS
23 June 2021

Migratory birds aid the redistribution of plants to new climates

Birds that travel long distances can disperse seeds far and wide. An assessment of the timing and direction of European bird migration reveals how these patterns might affect seed dispersal as the planet warms.

Barnabas H. Daru

 ORCID: http://orcid.org/0000-0002-2115-0257

0

Barnabas H. Daru

Barnabas H. Daru is in the Department of Life Sciences, Texas A&M University-Corpus Christi, Corpus Christi, Texas 78412, USA.

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The rapid pace of global warming and its effects on habitats raise the question of whether species are able to keep up so that they remain in suitable living conditions. Some animals can move fast to adjust to a swiftly changing climate. Plants, being less mobile, rely on means such as seed dispersal by animals, wind or water to move to new areas, but this redistribution typically occurs within one kilometre of the original plant1. Writing in Nature, González-Varo et al.2 shed light on the potential capacity of migratory birds to aid seed dispersal.When the climate in a plant’s usual range becomes hotter than it can tolerate, it must colonize new, cooler areas that might lie many kilometres away. It is not fully clear how plants distribute their seeds across great distances, let alone how they cross geographical barriers. One explanation for long-distance seed dispersal is through transport by migratory birds. Such birds ingest viable seeds when eating fruit (Fig. 1) and can move them tens or hundreds of kilometres outside the range of a plant species3. In this mode of dispersal, the seeds pass through the bird’s digestive tract unharmed4,5 and are deposited in faeces, which provides fertilizer that aids plant growth. In the case of European migratory birds, for example, the direction of seed dispersal will depend on whether the timing of fruit production coincides with a bird’s southward trip to warmer regions around the Equator, or northward to cooler regions. Many aspects of this process have been a mystery until now.

Figure 1 | A young blackcap bird (Sylvia atricapilla) eating elderberries.Credit: Getty

González-Varo and colleagues report how plants might be able to keep pace with rapid climate change through the help of migrating birds. The authors analysed the fruiting times of plants, patterns of bird migration and the interactions between fruit-eating birds and fleshy-fruited plants across Europe. Plants with fleshy fruits were chosen for this study because most of their seed transport is by migratory birds6, and because fleshy-fruited plants are an important component of the woody-plant community in Europe. The common approach until now has been to predict plant dispersal and colonization using models fitted to abiotic factors, such as the current climate. González-Varo et al. instead analysed an impressive data set of 949 different seed-dispersal interactions between bird and plant communities, together with data on entire fruiting times and migratory patterns of birds across Europe. The researchers also analysed DNA traces from bird faeces to identify the plants and birds responsible for seed dispersal.
Read the paper: Limited potential for bird migration to disperse plants to cooler latitudes
The authors hypothesized that the direction of seed migration depends on how the plants interact with migratory birds, the frequency of these interactions or the number of bird species that might transport seeds from each plant species. González-Varo and colleagues found that 86% of plant species studied might have seeds dispersed by birds during their southward trip towards drier and hotter equatorial regions in autumn, whereas only about one-third of the plant species might be dispersed by birds migrating north in spring. This dispersal trend was more pronounced in temperate plants than in the Mediterranean plant communities examined. These results are in general agreement with well-known patterns of fruiting times and bird migrations. For example, the fruit of most fleshy-fruited plants in Europe ripens at a time that coincides with when birds migrate south towards the Equator7.Perhaps the most striking feature of these inferred seed movements is the observation that 35% of plant species across European communities, which are closely related on the evolutionary tree (phylogenetically related), might benefit from long-distance dispersal by the northward journey of migratory birds. This particular subset of plants tends to fruit over a long period of time, or has fruits that persist over the winter. This means that the ability of plants to keep up with climate change could be shaped by their evolutionary history — implying that future plant communities in the Northern Hemisphere will probably come from plant species that are phylogenetically closely related and that have migrated from the south. Or, to put it another way, the overwhelming majority of plant species that are dispersed south towards drier and hotter regions at the Equator will probably be less able to keep pace with rapid climate change in their new locations than will the few ‘winners’ that are instead dispersed north to cooler climates. This has implications for understanding how plants will respond to climate change, and for assessing ecosystem functions and community assembly at higher levels of the food chain. However, for seeds of a given plant species, more evidence is needed to assess whether passing through the guts of birds affects germination success.To determine which birds might be responsible for the plant redistributions to cooler climates in the north, the authors categorized European bird migrants into Palaearctic (those that fly to southern Europe and northern Africa during their non-breeding season) and Afro-Palaearctic (those that winter in sub-Saharan Africa). Only a few common Palaearctic migrants, such as the blackcap (Sylvia atricapilla; Fig. 1) or blackbird (Turdus merula), provide most of this crucial dispersal service northwards to cooler regions across Europe. Because migratory birds are able to relocate a small, non-random subset of plants, this could well have a strong influence on the types of plant community that will form under climate-change conditions.
A bird’s migration decoded
A major problem, however, is that the role of these birds in dispersing seeds over long distances is already at risk from human pressures and environmental changes8. Understanding these large-scale seed-dispersal interactions offers a way for targeted conservation actions to protect the areas that are most vulnerable to climate change. This could include boosting protection efforts in and around the wintering grounds of migratory birds — locations that are already experiencing a rise in human pressures, such as illegal bird hunting.González-Varo and colleagues’ focus on seed dispersal across a Northern Hemisphere region means that, as with most ecological analyses, the results are dependent on scale, which can cause issues when interpreting data9. Because the Northern Hemisphere has more land area and steeper seasonal temperature gradients than the Southern Hemisphere does, seed-dispersal interactions might have different patterns from those occurring in the Southern Hemisphere or in aquatic systems.For example, seed-eating birds from the genus Quelea migrate from the Southern Hemisphere to spend the dry season in equatorial West Africa, then move southwards again when the rains arrive. Their arrival in southern Africa usually coincides with the end of the wet season in this region, when annual grass seeds are in abundance. It will be worth investigating whether migratory birds in the Southern Hemisphere also influence the redistribution of plant communities during global warming. Likewise, exploring the long-distance dispersal of seeds of aquatic plants, such as seagrasses10 by water birds, is another area for future research that might benefit from González-Varo and colleagues’ methods.This study provides a great example of how migratory birds might assist plant redistribution to new locations that would normally be difficult for them to reach on their own, and which might offer a suitable climate. As the planet warms, understanding how such biological mechanisms reorganize plant communities complements the information available from climate-projection models, which offer predictions of future species distributions.

doi: https://doi.org/10.1038/d41586-021-01547-1

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Competing Interests
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Network reconstruction from observational dataThe typical field study of plant–pollinator interactions involves recording instances of potential pollinators (such as insects) visiting plants within a prescribed observation area and over a prescribed period of time. We will refer to these records as visitation data. Network ecologists analyze visitation data by constructing networks of plant and pollinator species, where a connection between two species indicates that a plant-pollinator interaction exists between them.However, the meaning of edges in ecological networks is not always clear31. One popular way to transform visitation data into networks is to connect two species when they interact “enough”—say when a pollinator species is seen on the reproductive organ of a plant species a specified number of times—but in this case the precise meaning of an edge will depend on the details of the data collection and the choices made in the analysis. How many visits do we take as evidence of a plant–pollinator interaction? A single visit is probably not enough—it might well be an error or misobservation. Is two enough, or ten, or a hundred? And what about observations that were missed entirely? Other methods of analysis transform the data in different ways, for instance encoding them as weighted networks, possibly with some statistical processing along the way32. Even in this case, however, the edges still just count numbers of visits (perhaps transformed in some way), so the resulting networks are effectively histograms in disguise, recording only potential interactions rather than true biological connections.A more principled approach to network construction begins with a clear definition of what relationship (or relationships) a network’s edges encode33. We argue that network ecology often calls for a network of preferred interactions. In the context of plant-pollinator networks the edges of such a network indicate that pollinators preferentially visit certain plant species and they encode a variety of mechanisms that constrain species interactions, such as temporal or spatial uncoupling (i.e., species that do not co-occur in either time or space), constraints due to trait mismatches (e.g., proboscis size very different from corolla size), and physiological-biochemical constraints that prevent the interactions (e.g., chemical barriers). (One can regard preferred interactions as being the opposite of the “forbidden links” described in refs. 34,35,36). Preferred interactions are arguably the relevant ones for instance when analyzing the reaction of a network to abrupt changes: when one removes a plant species from a system, for example, the pollinators that prefer it will have to modify their behavior7,37,38. The interactions we consider are binary—either a species prefers another species or it doesn’t—so the network does not encode varying strengths of interaction.While the data gathered in a typical field study are certainly reflective of preferred interactions, they are, for many reasons, not perfect measurements of networks of preferred interactions13,17. First, there may be observational errors. While the observers performing the work are usually highly trained individuals, they may nonetheless make mistakes. They may confuse one species for another, which is particularly easy to do for small-bodied insects, or smaller species may be overlooked altogether. Observers may make correct observations but record them wrongly. And there will be statistical fluctuations in the number of visits of an insect species to a plant species over any finite time. For rare interactions there may even be no visits at all if we are unlucky. The insects themselves may also appear to make “mistakes” by visiting plants that they typically do not pollinate. These and other factors mean that the record of observed visits is an inherently untrustworthy guide to the true structure of the network of preferred interactions. Here we develop a statistical method for making estimates of network structure despite these limitations of the data.Model of plant–pollinator dataConsider a typical plant–pollinator study in which some number np of plant species, labeled by i = 1…np, and some number na of animal pollinator species, labeled by j = 1…na, are under observation for a set amount of time, producing a record of observed visits such that Mij is the number of times plant species i is visited by pollinator species j. Collectively the Mij can be regarded as a data matrix M with np rows and na columns. This is the input to our calculation.The unknown quantity, the thing we would like to understand, is the network of plant–pollinator interactions. We can think of this network as composed of two sets of nodes, one representing plant species and the other pollinator species, with connections or edges joining each pollinator to the plants it pollinates. In the language of network science this is a bipartite network, meaning that edges run only between nodes of unlike kinds—plants and pollinators—and never between two plants or two pollinators. Such a network can be represented by a second matrix B, called the incidence matrix, with the same size as the data matrix and elements Bij = 1 if plant i is preferentially visited by pollinator j and 0 otherwise.The question we would like to answer is this: What is the structure of the network, represented by B, given the data M? It is not straightforward to answer this question directly, but it is relatively easy to answer the reverse question. If we imagine that we know B, then we can say what the probability is that we make a specific set of observations M. And if we can do this then the methods of Bayesian inference allow us to invert the calculation and compute B from a knowledge of M and hence achieve our goal. The procedure is as follows.Consider a specific plant-pollinator species pair i, j. How many times do we expect to see j visit i if there is, or is not, a preferred interaction between i and j? The answer will depend on several factors. First, and most obviously, we expect the number of visits to be higher if j is in fact a pollinator of i. That is, we expect Mij to be larger if Bij = 1 than if Bij = 0. Second, we expect there to be more visits if there is greater sampling effort—for instance if the period of observation is longer or if the land area over which observations take place is larger15,16,26,27. Third, we expect to see more visits for more abundant plant and pollinator species than for less abundant ones, as demonstrated by several studies28,30. And fourth, as discussed above, we expect there to be some random variation in the number of visits, driven by fluctuations in individual behavior and the environment. These are the primary features that we incorporate into our model. It is possible to add others to handle specific situations (see ref. 39 and the Methods), but we focus on these four here.We translate these factors into a mathematical model of plant–pollinator interaction as follows. The random variations in the numbers of visits will follow a Poisson distribution for each plant–pollinator pair i, j, parameterized by a single number, the distribution mean μij, provided only that measurements are made sufficiently far apart to be independent (which under normal conditions they will be). We expect μij to depend on the factors discussed above and we introduce additional parameters to represent this dependence. First we introduce a parameter r to represent the change in the average number of visits when two species are connected (Bij = 1), versus when they are not (Bij = 0). We write the factor by which the number of visits is increased as 1 + r with r ≥ 0, so that r = 0 implies no increase and successively larger values of r give us larger increases. Second, we represent the effect of sampling effort by an overall constant C that multiplies the mean μij. The same constant is used for all i and j, since the same sampling effort is devoted to all plant–pollinator pairs. Third, we assume that the number of visits is proportional to the abundance of the relevant plant and pollinator species: twice as many pollinators of species j, for instance, will mean twice as many visits by that species, and similarly for the abundance of the plant species13. Thus the number of visits will be proportional to σiτj, for some parameters σi and τj representing the abundances of plant i and pollinator j, respectively, in suitable units (which we will determine shortly).Putting everything together, the mean number of observed visits to plant i by pollinator j is$${mu }_{ij}=C{sigma }_{i}{tau }_{j}(1+r{B}_{ij}),$$
(1)
and the probability of observing exactly Mij visits is drawn from a Poisson distribution with this mean:$$P({M}_{ij}| {mu }_{ij})=frac{{mu }_{ij}^{{M}_{ij}}}{{M}_{ij}!} {e}^{-{mu }_{ij}}.$$
(2)
This equation gives us the probability distribution of a single element Mij of the data matrix. Then, combining Eqs. (1) and (2), the data likelihood—the probability of the complete data matrix M—is given by the product over all species thus:$$P({boldsymbol{M}}| {boldsymbol{B}},theta )=mathop{prod}limits_{i,j}frac{{left[C{sigma }_{i}{tau }_{j}(1+r{B}_{ij})right]}^{{M}_{ij}}}{{M}_{ij}!} {e}^{-C{sigma }_{i}{tau }_{j}(1+r{B}_{ij})},$$
(3)
where θ is a shorthand collectively denoting all the parameters of the model: C, r, σ and τ. Our model is thus effectively a model of an entire network, rather than single interactions, in contrast with other recent approaches to the modeling of network data reliability17,18,32.There are two important details to note about this model. First, the definition in Eq. (1) does not completely determine C, σ, and τ because we can increase (or decrease) any of these parameters by a constant factor without changing the resulting value of μij if we simultaneously decrease (or increase) one or both of the others. In the language of statistics we say that the parameters are not “identifiable.” We can rectify this problem by fixing the normalization of the parameters in any convenient fashion. Here we do this by stipulating that σi and τj sum to one, thus:$$mathop{sum }limits_{i=1}^{{n}_{p}}{sigma }_{i}=mathop{sum }limits_{j=1}^{{n}_{a}}{tau }_{j}=1.$$
(4)
In effect, this makes σi and τj measures of relative abundance, quantifying the fraction of individual organisms that belong to each species, rather than the total number. (This definition differs from traditional estimates of pollinator abundance that define the abundance of a pollinator species in terms of its number of observed visits.) Second, there may be other species-level effects on the observed number of visits in addition to abundance, such as the propensity for observers to overlook small-bodied pollinators. There is, at least within the data used in this paper, no way to tell these effects from true variation in abundance—no way to tell for example if there are truly fewer individuals of a species or if they are just hard to see and hence less often observed. As a result, the abundance parameters in our model actually capture a combination of effects on observation frequency. This does not affect the accuracy of the model, which works just as well either way, but it does mean that we have to be cautious about interpreting the values of the parameters in terms of actual abundance. This point is discussed further in the applications below.Bayesian reconstructionThe likelihood of Eq. (3) tells us the probability of the data M given the network B and parameters θ. What we actually want to know is the probability of the network and parameters given the data, which we can calculate by applying Bayes’ rule in the form$$P({boldsymbol{B}},theta | {boldsymbol{M}})=frac{P({boldsymbol{M}}| {boldsymbol{B}},theta )P({boldsymbol{B}}| theta )P(theta )}{P({boldsymbol{M}})}.$$
(5)
This is the posterior probability that the network has structure B and parameter values θ given the observations that were made. There are three important parts to the expression: the likelihood P(M∣B, θ), the prior probability of the network P(B∣θ), and the prior probability of the parameters P(θ). The denominator P(M) we can ignore because it depends on the data alone and will be constant (and hence irrelevant for our calculations) once M is determined by the observations.Of the three non-constant parts, the first, the likelihood, we have already discussed—it is given by Eq. (3). For the prior on the network P(B∣θ) we make the conservative assumption—in the absence of any knowledge to the contrary—that all edges in the network are a priori equally likely. If we denote the probability of an edge by ρ, then the prior probability on the entire network is$$P({boldsymbol{B}}| theta )=mathop{prod}limits_{i,j}{(1-rho )}^{1-{B}_{ij}}{rho }^{{B}_{ij}}.$$
(6)
We consider ρ an additional parameter which is to be inferred from the data and which we will henceforth include, along with our other parameters, in the set θ.To complete Eq. (5), we also need to choose a prior P(θ) over the parameters. We expect there to be some limit on the value of r, which we impose using a minimally informative prior with finite mean (this distribution turns out to be the exponential distribution). For the remaining parameters we use uniform priors. With these choices, we then have everything we need to compute the posterior probability, Eq. (5).Once we have the posterior probability there are a number of things we can do with it. The simplest is just to maximize it with respect to the unknown quantities B and θ to find the most likely structure for the network and the most likely parameter values, given the data. This, however, misses an opportunity for more detailed inference and can moreover give misleading results. In most cases there will be more than one value of B and θ with high probability under Eq. (5): there may be a unique maximum of the probability, a most likely value, but there are often many other values that have nearly as high probability and offer plausible network structures competitive with the most likely one. To get the most complete picture of the structure of the network we should consider all these plausible structures.For example, if all plausible structures are similar to one another in their overall shape then we can be quite confident that this shape is reflective of the true preferred interactions between plant and pollinator species. If plausible structures are widely varying, however, then we have many different candidates for the true structure and our certainty about that structure is correspondingly lower. In other words, by considering the complete set of plausible structures we can not only make an estimate of the network structure but also say how confident we are in that estimate, in effect putting “error bars” on the network.How do we specify these errors bars in practice? One way is to place posterior probabilities on individual edges in the network. For example, when considering the edge connecting plant i and pollinator j, we would not ask “Is there an edge?” but rather “What is the probability that there is an edge?” Within the formulation outlined above, this probability is given by the average$$P({B}_{ij}=1| {boldsymbol{M}})=mathop{sum}limits_{{boldsymbol{B}}}int {B}_{ij}P({boldsymbol{B}},theta | {boldsymbol{M}})dtheta ,$$
(7)
where the sum runs over all possible incidence matrices and the integral over all parameter values. More generally we can compute the average of any function f(B, θ) of the matrix B and/or the parameters θ thus:$$leftlangle f({boldsymbol{B}},theta )rightrangle =mathop{sum}limits_{{boldsymbol{B}}}int f({boldsymbol{B}},theta ) P({boldsymbol{B}},theta | {boldsymbol{M}})dtheta .$$
(8)
Functions of the matrix and functions of the parameters can both be interesting—the matrix tells us about the structure of the network but the parameters, as we will see, can also reveal important information.Computing averages of the form (8) is unfortunately not an easy task. A closed-form expression appears out of reach and the brute-force approach of performing the sums and integrals numerically over all possible networks and parameters is computationally intractable in all but the most trivial of cases. The sum over B alone involves ({2}^{{n}_{p}{n}_{a}}) terms, which is normally a very large number.Instead therefore we use an efficient Monte Carlo sampling technique to approximate the answers. We generate a sample of network/parameter pairs (B1, θ1), …, (Bn, θn), where each pair appears with probability proportional to the posterior distribution of Eq. (5). Then we approximate the average of f(B, θ) as$$leftlangle f({boldsymbol{B}},theta )rightrangle simeq frac{1}{n}mathop{sum }limits_{i=1}^{n}f({{boldsymbol{B}}}_{i},{theta }_{i}).$$
(9)
Under very general conditions, this estimate will converge to the true value of the average asymptotically as the number of Monte Carlo samples n becomes large. Full details of the computations are given in Materials and Methods, and an extensive simulation study of the model is presented in Supplementary Note 1.Checking the modelInherent in the discussion so far is the assumption that the data can be well represented by our model. In other words, we are assuming there is at least one choice of the network B and parameters θ such that the model will generate data similar to what we see in the field. This assumption could be violated if our model is a poor one, but there is nothing in the method described above that would tell us so. To be fully confident in our results we need to be able not only to infer the network structure, but also to check whether that structure is a good match to the data. The Bayesian toolbox comes with a natural procedure for doing this. Given a set of high-probability values of B and θ generated by the method, we can use them in Eq. (3) to compute the likelihood P(M∣B, θ) of a data set M and then sample possible data sets from this probability distribution, in effect recreating data as they would appear if the model were in fact correct. We can then compare these data to the original field data to see if they are similar: if they are then our model has done a good job of capturing the structure in the data.In the parlance of Bayesian statistics this approach is known as a posterior–predictive assessment40. It amounts to calculating the probability$$P({widetilde{M}}_{ij}| {boldsymbol{M}})=mathop{sum}limits_{{boldsymbol{B}}}int P({widetilde{M}}_{ij}| {boldsymbol{B}},theta )P({boldsymbol{B}},theta | {boldsymbol{M}})dtheta$$
(10)
that pollinator species j makes ({widetilde{M}}_{ij}) visits to plant species i in artificial data sets generated by the model, averaged over many sets of values of B and θ. We can then use this probability to calculate the average value of ({widetilde{M}}_{ij}) thus:$$langle {widetilde{M}}_{ij}rangle =mathop{sum}limits_{{widetilde{M}}_{ij}}{widetilde{M}}_{ij} P({widetilde{M}}_{ij}| {boldsymbol{M}}).$$
(11)
The averages for all plant–pollinator pairs can be thought of as the elements of a matrix (langle widetilde{{boldsymbol{M}}}rangle), which we can then compare to the actual data matrix M, or alternatively we can calculate a residue ({boldsymbol{M}}-langle widetilde{{boldsymbol{M}}}rangle). If (langle widetilde{{boldsymbol{M}}}rangle) and M are approximately equal, or equivalently if the residue is small, then we consider the model a good one.To quantify the level of agreement between the fit and the data we can also compute the discrepancy40 between the artificial data and M as$${X}^{2}=mathop{sum}limits_{ij}frac{{({M}_{ij}-{langle widetilde{M}_{ij}rangle })}^{2}}{{langle widetilde{M}_{ij}rangle }}.$$
(12)
Under the hypothesis that the model is correct, X2 follows a chi-squared distribution with np × na degrees of freedom40. A good fit between model and data is signified by a value of X2 that is much smaller than its expectation value of np × na. Note that the calculation of (P({widetilde{M}}_{ij}| {boldsymbol{M}})) in Eq. (10) is of the same form as the one in Eq. (8), with (f({boldsymbol{B}},theta )=P({widetilde{M}}_{ij}| {boldsymbol{B}},theta )), which means we can calculate (P({widetilde{M}}_{ij}| {boldsymbol{M}})) in the same way we calculate other average quantities, using Monte Carlo sampling and Eq. (9).Application to visitation data setsWell-sampled dataTo demonstrate how the method works in practice, we first consider a large data set of plant–pollinator interactions gathered by Kaiser-Bunbury and collaborators41 at a set of study sites on the island of Mahé in the Seychelles. The data describe the interactions of plant and pollinator species observed over a period of eight months across eight different sites on the island. The data also include measurements of floral abundances for all observation periods and all sites. Our method for inferring network structure does not make use of the abundance measurements, but we discuss them briefly at the end of this section.The study by Kaiser-Bunbury et al. focused particularly on the role of exotic plant species in the ecosystem and on whether restoring a site by removing exotic species would significantly impact the resilience and function of the plant–pollinator network. To help address these questions, half of the sites in the study were restored in this way while the rest were left unrestored as a control group.As an illustration of our method we apply it to data from one of the restored sites, as observed over the course of a single month in December 2012 (the smallest time interval for which data were available). We pick the site named “Trois-Frères” because it is relatively small but also well sampled. Our calculation then proceeds as shown in Fig. 1. There were 8 plant and 21 pollinator species observed at the site during the month, giving us an 8 × 21 data matrix M as shown in Fig. 1a. (Following common convention, the plots of matrices in this paper are drawn with rows and columns ordered by decreasing numbers of observed interactions, so that the largest elements of the data matrix—the darkest squares—are in the top and left of the plot.)Fig. 1: Illustration of the method of this paper applied to data from the study of Kaiser-Bunbury et al.41.a We start with a data matrix M that records the number of interactions between each plant species and pollinator species. Species pairs that are never observed to interact (Mij = 0) are shown in white. b We then draw 2000 samples from the distribution of Eq. (5), four of which are shown in the figure. Each sample consists of a binary incidence matrix B, values for the relative abundances σ and τ (shown as the orange and blue bar plots, respectively), and values for the parameters C, r, and ρ (not shown). c We combine the samples using Eqs. (7)–(9) to give an estimate of the probability of each edge in the network and the complete parameter set θ. For the data set studied here our estimates of the expected values of the parameters C, r, and ρ are 〈C〉 = 20.2, 〈r〉 = 45.9, and 〈ρ〉 = 0.244.Full size imageNow we use our Monte Carlo procedure to draw 2000 sets of incidence matrices B and parameters θ from the posterior distribution of Eq. (5) (Fig. 1b). These samples vary in their structure: some edges, like the one connecting the plant N. vanhoutteanum and the pollinator A. mellifera, are present in nearly all samples, while others, like the one between M. sechellarum and A. mellifera, appear only a small fraction of the time. Some others never occur at all. Averaging over these sampled networks we can estimate the probability, Eq. (7), that each connection exists in the network of preferred interactions between plant and animal species—see Fig. 1c. Some connections have high probability, close to 1, meaning that we have a high degree of confidence that they exist. Others have probability close to 0, meaning we have a high degree of confidence that they do not exist. And some have intermediate probabilities, meaning we are uncertain about them (such as the M. sechellarum–A. mellifera connection, which has probability around 0.45). In the latter case the method is telling us that the data are not sufficient to reach a firm conclusion about these connections. Indeed, if we compare with the original data matrix M in Fig. 1a, we find that most of the uncertain connections are ones for which we have very few observations, relative to the total number of observations for these species—say Mij = 1 or 2 for species with dozens of total observations overall.As we have mentioned, we also need to check whether the model is a good fit to the data by performing a posterior–predictive test. Figure 2 shows the results of this test. The main plot in the figure compares the values of the 40 largest elements of the original data matrix M with the corresponding elements of the generated matrix (widetilde{{boldsymbol{M}}}). In each case, the original value is well within one standard deviation of the average value generated by the test, confirming the accuracy of the model. The inset of the figure shows the residue matrix ({boldsymbol{M}}-widetilde{{boldsymbol{M}}}), which reveals no systematic bias unaccounted for by the model. The discrepancy X2 of Eq. (12) takes the value 26.94 in this case, well below the expected value of npna = 168, which indicates that the good fit is not a statistical fluke.Fig. 2: Results of a posterior–predictive test on the data matrix M for the example data set analyzed in Fig. 1.The main plot shows the error on the 40 largest entries of M, while the inset shows the residue matrix ({boldsymbol{M}}-langle tilde{{boldsymbol{M}}}rangle). Because the actual data M are well within one standard deviation of the posterior–predictive mean, the test confirms that the model is a good fit in this case. Error bars correspond to one standard deviation and are computed with n = 2000 samples from the posterior distribution.Full size imageIn addition to inferring the structure of the network itself, our method allows us to estimate many other quantities from the data. There are two primary methods by which we can do this. One is to look at the values of the fitted model parameters, which represent quantities such as the preference r and species abundances σ, τ. The other is to compute averages of quantities that depend on the network structure or the parameters (or both) from Eq. (9).As an example of the former approach, consider the parameter ρ, which represents the average probability of an edge, also known as the connectance of the network. Figure 3a shows the distribution of values of this quantity over our set of Monte Carlo samples, and neatly summarizes our overall certainty about the presence or absence of edges. If we were certain about all edges in the network, then ρ would take only a single value and the distribution would be narrowly peaked. The distribution we observe, however, is somewhat broadened, indicating significant uncertainty. The most likely value of ρ, the peak of the distribution, turns out to be quite close to the value one would arrive at if one were simply to assume that every pair of species that interacts even once is connected in the network. This does not mean, however, that one could make this assumption and get good results. As we show below, the network one would derive by doing so would be badly in error in other ways.Fig. 3: Analyses that can be performed using samples from the posterior distribution of Eq. (5).a Distribution of the connectance ρ. Connectance values for binary networks obtained by thresholding the data matrix at Mij  > 0 and Mij ≥ 5 are shown as vertical lines for reference. b Distribution of the preference parameter r. The mean value of r is 〈r〉 = 45.9 and its mode close to 40, but individual values as high as 100 are possible. c Distribution of the nestedness measure NODF. Values obtained by thresholding the data matrix at Mij  > 0 and Mij  > 1 are shown for reference. d Measured and estimated abundances for each of the plant species (R2 = 0.54).Full size imageFigure 3b shows the distribution of another of the model parameters, the parameter r, which measures the extent to which pollinators prefer the plants they normally pollinate over the ones they do not. For this particular data set the most likely value of r is around 40, meaning that pollinators visit their preferred plant species about 40 times more often than non-preferred ones, indicating all other things being equal, an impressive level of selectivity on the part of the pollinators.For the calculation of more complicated network properties we can perform an average over the value of any function f(B, θ), as long as there is an algorithm to compute it. As an example, Fig. 3c shows a calculation of the quantity known as “Nestedness based on Overlap and Decreasing Fill” (NODF), a measure of the nestedness property discussed in the introduction. This quantity measures the extent to which specialist species—those with relatively few interactions—tend to interact with a subset of the partners of generalist species42. While it is complicated to compute NODF analytically, due to the fact that one must order the species by degrees22, it is straightforward to calculate it within our framework: we simply calculate the value for each sampled network B and plot the resulting distribution. Interestingly, the most likely value of NODF is significantly different from the one we would calculate had we assumed, as discussed above, that a single interaction is sufficient to consider two species connected. On the contrary, we find that the system is almost certainly more nested than this simple analysis would conclude.In Fig. 3d, we compare the values of our estimated floral abundance parameters σ to the measured abundances reported by Kaiser-Bunbury et al.41. These parameters are not measures of abundance in the usual sense, because they combine actual abundance (quantity or density) with other characteristics such as ease of observation. We do find a correlation between the estimated and observed abundances, but it is relatively weak (R2 = 0.54), signaling significant disagreement, on which we elaborate in the discussion section.Undersampled dataAs we have pointed out, the connections in the network about which we are most uncertain tend to be ones that are undersampled, i.e., those for which we have only a small amount of data. In an ideal world we could address this problem by taking more data, but it is rare that we have the opportunity to do this. More commonly the data have already been gathered and our task is to produce the best results we can with those data. There are nonetheless some remedies open to us, such as aggregating data over different geographical areas or time windows. In Fig. 4 we compare the edge probabilities estimated from data recorded individually at the four “restored” sites in the Mahé study during October 2012 to the edge probabilities we obtain when we aggregate these observations into a single data matrix and only then estimate the network. (We use restored sites observed during the same month because they are likely to be ecologically similar, meaning the data are measuring approximately the same system.) Comparison of the two distributions shows—as we would hope—that there are fewer uncertain edges in the aggregated network than in its disaggregated parts, i.e., there are fewer edges with probabilities in the middle of the distribution and more with probabilities close to zero or one.Fig. 4: Illustration of the effect of data aggregation on edge uncertainty.a Histogram of the edge probabilities P(Bij = 1∣M) for the four restored sites in the Mahé study as observed in October 2012 and analyzed individually. b Equivalent histogram after aggregating the data over the sites and then estimating a single network from the resulting data matrix. The horizontal lines, both drawn at fifty observations—are added merely as a guide to the eye. Note how the upper histogram has more mass near the middle of the plot, while the lower one has most of its mass close to probability zero or one, indicating greater certainty in the positions of the edges in the aggregated data.Full size imageIn other cases neither aggregation nor gathering more data is possible, for instance when reanalyzing a data set already collected by others or already maximally aggregated. Such data sets record the results of observational studies that are already over, and may contain too few observations, but our approach still allows us to perform rigorous inference in these circumstances.For instance, Jordano et al.43 used dozens of existing plant-pollinator and plant-frugivore data sets to argue that the degree distributions of mutualistic networks have a long tail, but this conclusion is undermined by issues with undersampling. As an example, one of the data sets they studied, originally gathered by Inouye and Pyke44, records 1314 individual interactions over a period of 3 months in Kosciusko National Park, Australia, between 40 plants and 85 pollinator species, which works out to an average of 0.386 unique observations per species pair. Is this sampling effort sufficient to establish edges with certainty? As a point of reference, the data analyzed in Fig. 1 comprises 201 observations between 8 plants and 21 pollinators species for an average of 1.196 observations per pair of species, and the aggregated data of Fig. 4 contain 1.420 observations for every pair. Nonetheless, there is uncertainty about some of the connections in these reconstructed networks; this suggests that the network of Inouye and Pyke, with less than a third as much data per species pair, will contain significant uncertainty.Even so, our method allows us to make inferences about this network. In Fig. 5, we show estimates of the degree distributions of both plant and pollinator nodes in the network obtained from the posterior distribution P(B∣M), along with naive estimates calculated by thresholding the (undersampled) data as in the study by Jordano et al.43. As the figure shows, the results derived from the two approaches are very different. The thresholded degree distributions were classified as scale-free by Jordano et al., but this classification no longer holds once we account for the issues with the data; the inferred degree distributions are in this case well-modeled as Poisson distributions of means 5.53 and 2.60 for plants and pollinators respectively and the power-law form is a poor fit. On the other hand, the abundance parameters of the model, shown in Fig. 5, do appear to have a broad distribution, an interesting finding that calls for a rethinking of the relationship between abundances and degree distributions. It is generally thought that interactions will tend to be evenly distributed under an even distribution of abundance13 but here the opposite seems to be true.Fig. 5: Distributions of species-level parameters for a network of plants and pollinators in Kosciusko National Park, Australia, from the study by Inouye and Pyke44.a Thresholded degree distributions calculated by connecting species i and j with an edge if Mij  > 0. Inferred degree distributions are calculated using the method of this paper, averaging the fraction pk of nodes with a given degree k over n = 2000 Monte Carlo samples. b Inferred distributions of abundances σ and τ, calculated as a histogram over n = 2000 Monte Carlo samples of the abundance parameters of the fitted model. Error bars correspond to one standard deviation in all cases.Full size image More

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Bacterial strain and growth conditionsWe used Pseudomonas protegens (formerly Pseudomonas fluorescens)55 CHA0 as a model strain, which was initially isolated from tobacco roots56. The strain was chromosomally tagged with GFP and a kanamycin resistance cassette to enable specific tracking of the strain and detection of contaminations19. This bacterium has the genetic potential to produce various bioactive metabolites, including the plant hormone indole-3-acetic acid (IAA), antimicrobial compounds and lytic enzymes25. Prior to the experiment, bacteria were grown for 48 h on a King′s medium B57 (KB) agar plate supplemented with 50 µg ml−1 kanamycin, a single colony was randomly picked and grown for 12 h in KB at 28 °C with agitation. The cell culture was then washed for three times in 10 mM MgSO4 and adjusted to 107 cells ml−1 and used as inoculant for all plants. This inoculant was also stored at −80 °C as frozen ancestral stock, from which ʻAncestorʼ isolates were picked in later experiments.Host plant and growth conditionsWe used Arabidopsis thaliana ecotype Col-0 as a model host plant. Surface-sterilized seeds were first sown in Petri dishes with agar-solidified (1.5% agar (w/v)) modified Hoagland′s medium: (KNO3 (3 mM), MgSO4 (0.5 mM), CaCl2 (1.5 mM), K2SO4 (1.5 mM), NaH2PO4 (1.5 mM), H3BO3 (25 μM), MnSO4 (1 μM), ZnSO4 (0.5 μM), (NH4)6Mo7O24 (0.05 μM), CuSO4 (0.3 μM), MES (2.5 mM) and 50 μM Fe(III)EDTA, pH = 5.8) and stratified for 2 days at 4 °C after Petri dishes were positioned vertically and transferred to a growth chamber (20 °C, 10 h light/14 h dark, light intensity 100 μmol m−2 s−1). After 2 weeks of incubation, two seedlings were transferred to closed and sterile ECO2 boxes (http://www.eco2box.com/ov80xxl_nl.htm) for selection experiment. The ECO2 boxes were filled with 260 g of dry, carbon-free silver sand that was previously washed with MilliQ water to remove dissolvable chemical elements and heated to 550 °C for 24 h to remove remaining organic material. Prior to transplantation the sand was amended with 13 ml of modified Hoagland medium.Design of the selection experimentThe selection experiment was conducted in a gnotobiotic system to remove confounding effects that may emerge as a result of competitive interactions with other microorganisms, and to place the focus on plant-mediated selective pressures. Moreover, we allowed only the bacteria to evolve during the experiment and used new clonal plants at every bacterial transfer. We set up five independent plant–bacterium replicate lines, which were grown for six independent plant growth cycles (see Fig. S1 for an overview of the experimental design). The experiment was started by inoculating 106 cells of the stock P. protegens CHA0 culture (from here on abbreviated as ʻancestorʼ) into the rhizosphere of 2-week-old A. thaliana seedlings growing in sterile silver sand within ECO2 boxes (two plants per replicate selection line). Inoculated plants were then grown for 4 weeks (20 °C, 10 h light/14 h dark, light intensity 100 μmol m−2 s−1) after which the plant growth cycle was terminated and root-associated bacteria were harvested by placing the roots of both plants into a 1.5 ml Eppendorf tubes filled with 1 ml 10 mM MgSO4 and two glass beads. Rhizosphere bacteria were suspended into the liquid using a TissueLyser II at a frequency of 20 s−1 for 1 min after which bacterial cell densities were determined using flow cytometry (BD Accuri™ C6 Plus, thresholds for FSC: 2000, SSC: 8000). After this, 106 cells were inoculated to the rhizosphere of new A. thaliana plants to initiate the next plant growth cycle. Possible contaminations were checked by plating the suspension on 3 g l−1 tryptic soy agar (TSA) plates and it was verified that all colonies carried the GFP marker gene, as observed under UV light.Bacterial life-history traits measurementsIndividual bacterial colonies were isolated from all replicate plant selection lines for life-history measurements at the end of the second, fourth and sixth plant growth cycle by dilution plating the rhizosphere suspension on 3 g l−1 TSA plates. After incubation at 28 °C for 24 h, 16 colonies were randomly picked from each replicate selection lines resulting in a total of 240 evolved and 16 ancestral colonies. All these colonies were characterized for a set of key bacterial life-history traits representative of growth, stress resistance and traits linked with plant–microbe interactions.

a.

Bacterial growth yield in KB medium
All the bacterial isolates were grown in 96-well plates with 160 µl 1/3 strength liquid KB, at 20 °C without shaking. Bacterial yield was determined as the maximum optical density at 600 nm after 3 days of growth using a spectrophotometer (SPECTROstar Nano).

b.

Bacterial stress resistance
We measured bacterial resistance to a range of different stresses using various 96-well microplate assays. Abiotic stress resistance was determined by growing bacteria in 160 µl of 1 g l−1 TSB containing 0.0025% H2O2 (oxidative stress), 15% polyethylene glycol (PEG)−6000 (water potential stress) or 2% NaCl (salt stress). We used resistance to antibiotics commonly produced by rhizosphere microorganisms as indicator of biotic stress resistance. Antibiotic resistance was tested in 160 µl of 1 g l−1 TSB supplemented with 1 µg ml−1 streptomycin, 1 µg ml−1 tetracycline, or 5 µg ml−1 penicillin, respectively. Bacterial growth were determined after 3 days of growth at 20 °C without shaking as optical density at 600 nm.

c.

Traits linked with plant–microbe interactions
P. protegens CHA0 harbours several traits that are linked to plant growth including production of antibiotics and plant hormones. To assess these traits, we grew each bacterial colony in 96-well plates containing 160 µl of 1/3 strength liquid KB per well at 20 °C without shaking for 72 h. Cell-free supernatants were obtained by filter sterilization (0.22 µm) using Multiscreen HTS 96-well filtration plates (1000 × g, 30 min), which were used to measure the production of the plant hormone auxin (Indole-3-acetic acid (IAA)), iron-chelating siderophores and proteolytic activity. Furthermore, we also measured antifungal and antibacterial activity of all colonies.

IAA detectionThe production of the plant hormone auxin was determined with a colorimetric test58. Briefly, 30 µl P. protegens CHA0 cell-free filtrate was incubated with 30 µl R1 reagent (12 g l−1 FeCl3, 7.9 M H2SO4) for 12 h in the dark and optical density read at 530 nm of the colorimetric complex was used as a measurement of IAA concentration.Siderophore activityIron-chelating ability was measured as a proxy for siderophore production59. To this end, 100 µl of P. protegens CHA0 cell-free filtrate was mixed with 100 µl of modified CAS solution (with 0.15 mM FeCl3) and optical density read at 630 nm after 3 h of incubation was used as a proxy of siderophore production. The iron-chelating ability was calculated based on the standard curve based on modified CAS assay solution with a range of iron concentration (0, 0.0015, 0.003, 0.006, 0.009, 0.012, 0.015 mM FeCl3).Proteolytic activityThe proteolytic activity assay we used was adapted from Smeltzer et al.60. Briefly, 15 µl of P. protegens CHA0 cell-free filtrate was incubated with 25 µl of azocasein (2% w/v in 50 mM Tris-HCl pH 8.0) at 40 °C for 24 h. One hundred and twenty-five of 10% w/v cold trichloroacetic acid (TCA) was added to precipitate superfluous azocasein, and then 100 µl supernatant was neutralized with 100 µl of 1 M NaOH after centrifugation at 5000g for 30 min. Optical density read at 440 nm was used as a proxy of exoprotease activity.Tryptophan side chain oxidase (TSO) activityTSO activity, an indicator of quorum sensing activity in P. protegens CHA0, was measured based on an modified established colorimetric assay61: Three-day-old bacterial cultures grown in 1/3 strength liquid KB were mixed at a 1:1 ratio with a reagent solution (5 g l−1 SDS, 37.6 g l−1 glycine 2.04 l−1 g tryptophan, pH 3.0) and TSO activity was measured as optical density at 600 nm after overnight incubation.Biofilm formationWe quantified bacterial biofilm formation using a standard protocol62. Briefly, bacteria were grown at 20 °C for 72 h in 160 µl 1 g l−1 TSB in a 96-well microtiter plate with TSP lid (TSP, NUNC, Roskilde, Denmark). Planktonic cells were removed by immersing the lid with pegs three times in phosphate-buffered saline solution (PBS). Subsequently, the biofilm on the pegs was stained for 20 min in 160 µl 1% crystal violet solution. Pegs were washed five times in PBS after which the crystal violet was extracted for 20 min from the biofilm in a new 96-well microtiter plate containing 200 µl 96% ethanol per well. Biofilm formation was defined as the optical density at 590 nm of the ethanol extracted crystal violet63.Inhibition of other microorganismsAntimicrobial activity was defined as the relative growth of the target organism in P. protegens supernatant compared to the control treatment. Antifungal activity of the cell-free supernatant was assessed against the ascomycete Verticillium dahliae. The fungus was grown on potato dextrose agar at 28 °C for 4 days, after which plugs of fungal mycelium were incubated in potato dextrose broth medium at 28 °C and gentle shaking for 5 days. Fungal spores were collected by filtering out the mycelium from this culture over glass wool. Subsequently, spores were washed and resuspended in water and the OD595 of the suspension was adjusted to 1. Five microlitres of this spore suspension was then inoculated with 15 µl P. protegens CHA0 cell-free filtrate and incubated in 160 µl of 1 g l−1 PDB medium for 2 days at 20 °C in 96-well plates. Fungal growth was measured as optical density at 595 nm after 2 days of growth and contrasted with the growth in the control treatment (PDB medium without P. protegens supernatant). Antibacterial activity was determined using the plant pathogen Ralstonia solanacearum as a target organism. R. solanacearum was grown in 160 µl of 1 g l−1 TSB medium supplemented with 15 μl of P. protegens CHA0 cell-free filtrate or 15 µl of 1/3 strength liquid KB as a control for 2 days at 20 °C. R. solanacearum growth was measured as optical density at 600 nm.Determining changes in P. protegens CHA0 interactions with A. thaliana after the selection experimentBased on the life-history trait measurements, five distinct bacterial phenotypes were identified using K-means clustering analysis (Fig. S2). In order to assess whether phenotypic changes reflected shifts in the strength and type of plant–bacterium interaction, we chose five isolates from each bacterial phenotype group representing each replicate selection line and five ancestral isolates for further measurements (a total of 30 isolates, Table S2).Effects of ancestor and evolved bacteria on plant performanceFor each isolate we measured root colonizing ability and impact on plant performance. All 30 bacterial isolates were incubated overnight in 1/3 KB strength liquid at 20 °C. The culture was centrifuged twice for 5 min at 5000 × g and the pellet was washed and finally resuspended in 10 mM MgSO4. The resulting suspension was adjusted to an OD600 of 0.01 for each strain64. Ten microlitres of the bacterial suspension (or 10 mM MgSO4 as a control) was applied to the roots of three 10-day old sterile Arabidopsis thaliana Col-0 seedlings (excluding 2 days of stratification at 4 °C) grown on vertically positioned Petri dishes with agar-solidified (1.5% agar (w/v)) modified Hoagland′s medium (n = 3 biological plant replicates, each containing 3 seedlings). Plants were grown for 14 days before harvesting. Plants were photographed before and 14 days after bacterial inoculation.Bacterial effects on plant health were quantified as leaf ʻgreennessʼ as the presence of ancestral strain was observed to lead to bleaching and loss of chlorophyll in A. thaliana leaves. The ʻgreennessʼ was quantified from photographs by measuring the number of green pixels. To this end, photographs were first transformed in batch using Adobe Photoshop 2021 by sequentially selecting only green areas followed by thresholding balancing green tissue over background noise (Level 80). This resulted in black-and-white images for further analysis, and the mean number of white pixels per fixed-sized region-of-interest of the aboveground tissue was subsequently determined as ʻgreennessʼ using ImageJ (version 1.50i). The numbers of lateral roots and the primary root length were also measured using ImageJ. The root morphology data measured at the end of the experiment was normalized with the data collected at the time of inoculation for each individual seedling.To determine shoot biomass, the rosette of each plant was separated from the root system with a razor blade and weighted. The roots were placed into a pre-weighted 1.5 ml Eppendorf tubes to quantify the root biomass. To determine the bacterial abundance per plant, these tubes were subsequently filled with 1 ml 10 mM MgSO4 buffer solution and two glass beads. The rhizosphere bacteria were suspended into the buffer solution using a TissueLyser II at a frequency of 20 s−1 for 1 min after which bacterial densities were determined using flow cytometry (BD Accuri™ C6 Plus, thresholds for FSC: 2000, SSC: 8000). Shoot biomass, root biomass, root length and number of lateral roots were used in a principal component analysis (PCA) to calculate an overall impact of the bacteria on plant performance (Fig. 2e). The first principal component (PC1) explained 79.9% of the variation and was normalized against the control treatment to be used as a proxy of ʻPlant performanceʼ in which positive values reflect plant growth promotion and negative values plant growth inhibition.Root derived carbon source utilizationTo measure changes in bacterial growth on potential root derived carbon sources, we measured the growth of all 256 isolates using modified Ornston and Stanier (OS) minimal medium65 supplemented with single carbon sources at a final concentration of 0.5 g l−1 in 96-well plates containing 160 µl carbon supplemented OS medium per well. The following carbon sources were selected based on their relatively high abundance in Arabidopsis root exudates21: alanine, arabinose, butyrolactam, fructose, galactose, glucose, glycerol, glycine, lactic acid, putrescine, serine, succinic acid, threonine and valine. Bacterial growth was determined by measuring optical density at 600 nm after 3 days incubation at 20 °C.GUS histochemical staining assay and bacterial growth under scopoletin stressTo investigate effects of the ancestor and evolved strains of P. protegens CHA0 on expression of MYB72, we applied a GUS histochemical staining assay to the 30 selected isolates (Table S2). MYB72 is a transcription factor involved in production of the coumarin scopoletin in Arabidopsis roots and specific rhizobacteria can upregulate expression of MYB72 in the roots66. Scopoletin is an iron-mobilizing phenolic compound with selective antimicrobial activity22. Seedlings of the A. thaliana MYB72pro:GFP-GUS24 reporter line were prepared as described above. Seven-day-old seedlings were inoculated directly below the hypocotyls with 10 μl of a bacterial suspension (OD660 = 0.1)24. At 2 days after inoculation, the roots were separated from the shoots and washed in MilliQ water (Milliport Corp., Bedford, MA) to remove all the adhered bacteria. GUS staining of the roots was performed in 12-well microtiter plates where each well contained roots of 5–6 seedlings and 1 ml of freshly prepared GUS substrate solution (50 mM sodium phosphate with a pH at 7, 10 mM EDTA, 0.5 mM K4[Fe(CN)6], 0.5 mM K3[Fe(CN)6], 0.5 mM X-Gluc, and 0.01% Silwet L-77)67. Plates were incubated in the dark at room temperature for 16 h. The roots were fixed overnight in 1 ml ethanol:acetic acid (3:1 v/v) solution at 4 °C and transferred to 75% ethanol. Then the pictures of each microtiter plates were taken, and GUS activity was quantified by counting the number of blue pixels in each well of the microtiter plates using image analysis in ImageJ (version 1.52t). To assess the effects of scopoletin on ancestral and evolved P. protegens CHA0 isolates, we applied a sensitivity assay to the 30 selected isolates (Table S2). In brief, growth of bacterial isolates was measured in 1 g l−1 TSB medium (160 µl) supplemented with scopoletin at final concentrations of 0 µM (control), 500 µM, 1000 µM, and 2 mM using optical density at 600 nm after 72 h incubation at 20 °C without shaking in 96-well microtiter plates. Maximal effect (Emax) of scopoletin was calculated via R package ʻGRmetricsʼ68 as an indication of scopoletin tolerance.Whole-genome sequencingAll 30 isolated phenotypes were whole genome sequenced to identify possible mutations and affected genes. To this end, isolates were cultured overnight at 28 °C in 1/3 strength liquid KB. Chromosomal DNA was isolated from each culture using the GenElute™ Bacterial Genomic DNA Kit Protocol (NA2100). DNA samples were sheared on a Covaris E-220 Focused-ultrasonicator and sheared DNA was then used to prepare Illumina sequencing libraries with the NEBNext® Ultra™ DNA Library Prep Kit (New England Biolabs. France) and the NEBNext® Multiplex Oligos for Illumina® (96 Index Primers). The final libraries were sequenced in multiplex on the NextSeq 500 platform (2 × 75 bp paired-end) by the Utrecht Sequencing Facility (http://www.useq.nl) yielding between 1.0 and 6.4 million reads per sample equivalent to ~10–70-fold coverage (based on comparison with the original 6.8 Mbp reference genome NCBI GenBank: CP003190.1).Variant calling analysisWe first constructed an updated reference genome of P. protegens CHA0, carrying the GFP marker gene on its chromosome, from the ancestral strain using the A5 pipeline with default parameters69. The input dataset for this sample consisted of 3,1M reads and totals an approximate 34-fold coverage. The size of the updated reference genome is 6.8 Mbp, with a G + C content of 63.4%, and it comprises 80 scaffolds, with a N50 value of 343 kbp. We subsequently used PROKKA70 (version 1.12; https://github.com/tseemann/prokka) for full annotation of the updated reference genome, and this resulted in the identification of 6147 genes. The updated genome is deposited in NCBI GenBank with following reference: RCSR00000000.1.Having established the ancestral genome sequence, we subsequently used Snippy (version 3.2-dev; https://github.com/tseemann/snippy) to identify and functionally annotate single-nucleotide polymorphisms and small insertions and deletions (indels) for each individual strain. In addition, we investigated the breadth of coverage for each gene per sample with BedTools71 to identify genes with large insertions or deletions. An overview of the polymorphisms is shown in Supplementary Table S3. Raw sequencing data for this study are deposited at the NCBI database under BioProject PRJNA473919.Relative competitive fitness of gac mutants measured in vivo and in vitroThe relative competitive fitness of selected gac mutants was measured in direct competition with their direct ancestors both in vivo in the rhizosphere of A. thaliana and in vitro in different standard culture media. Relative fitness was measured as deviation from initial 1:1 ratio of bacterial clone pairs based on PCR-based high-resolution melting profile (RQ-HRM) analysis. Three pairs of isolates were selected: (A) evolved gacA ID 242 (genotype oafAY335X ∙ RS17350A77A.fsX14 ∙ gacAD49Y) and its direct ancestral genotype 133 (genotype oafAY335X ∙ RS17350A77A.fsX14) from evolutionary line 1; (B) evolved gacA ID 220 (genotype galEV32M ∙ accCE413K ∙ gacAD54Y) and its direct ancestral genotype 28 (genotype galEV32M ∙ accCE413K) from line 2; (c) evolved gacS ID 222 (genotype oafAK338S.fsX18 ∙ gacSG27D) and its direct ancestral genotype 66 (genotype oafAK338S.fsX18) from line 3. Bacterial isolates were first grown overnight in KB medium at 28 °C, centrifuged at 5000g for 10 min and the pellet resuspended in 10 mM MgSO4. This washing procedure was repeated twice. The resulting bacterial suspensions were diluted to OD600 = 0.05. The initial inoculum for the competition assays was then generated by mixing equal volumes of evolved and ancestral competitors in a ratio of 1:1.Measuring competitive fitness in A. thaliana rhizosphereThis assay was performed on the roots of 10-day old A. thaliana seedlings grown on full strength Hoagland agar plates, which were prepared as described earlier. Twenty microlitres of the initial inoculum, containing a total of 106 bacterial cells, was inoculated on to the root–shoot junction of each seedling. After 14 days of growth, bacterial populations were isolated from the roots and stored at −80 °C in 42.5% glycerol for relative abundance measurements.Measuring competitive fitness in culture mediaCompetition assays were also performed in three commonly used nutrient-rich growth media: KB, LB and TSB. KB contained 20 g proteose peptone, 1.5 g MgSO4.7H2O, 1.2 g KH2PO4 and 10 g glycerol per litre and the pH was adjusted to 7.3 ± 0.2. TSB contained 30 g tryptic soy broth per litre and pH was adjusted to 7.3 ± 0.2. LB contained 10 g peptone, 5 g yeast extract and 5 g NaCl per litre. Twenty microlitres inoculum of competing strains, containing about 106 bacterial cells, were added into wells containing 140 μl fresh medium in a 96-well plate. The microplates were incubated at 28 °C without shaking for 48 after 80 μl sample was harvested and stored at −80 °C in 42.5% glycerol from each well for relative abundance measurements.RQ-HRM assay for quantifying changes in genotype frequencies after competitionWe used a high-resolution melting (HRM) curve profile assay with integrated LunaProbes to quantify the ratio of mutant to wild-type genotypes72,73,74. The probes and primers used in this study are listed in Table S4. Primers were designed using Primer3. Probes were designed with the single-nucleotide polymorphism (SNP) located in the middle of the sequence, and the 3′ end was blocked by carbon spacer C3. The primer asymmetry was set to 2:1 (excess primer: limiting primer) in all cases. Pre-PCR was performed in a 10-μl reaction system, with 0.25 μM excess primer, 0.125 μM limiting primer, 0.25 μM probe, 0.5 μl bacterial sample culture (100-fold diluted saved sample, OD600 is about 0.01), 1× LightScanner Master Mix (BioFire Defense). DMSO with the final concentration 5% was supplemented in all reactions to ensure the targeted melting domains are within the detection limit of the LightScanner (Idaho Technology Inc.). Finally, MQ water was used to supplement up to 10 μl. A 96-well black microtiter plate with white wells was used to minimize background fluorescence. Before amplification, 25 μl mineral oil was loaded in each well to prevent evaporation, and the plate was covered with a foil seal to prevent the degradation of fluorescent molecules. Amplification was initiated by a holding at 95 °C for 3 min, followed by 55 cycles of denaturation at 95 °C for 30 s, annealing at 60 °C for 30 s and extension at 72 °C for 30 s and then kept at 72 °C for 10 min. After amplification, samples were heated in a ThermalCycler (Bio-Rad) shortly to 95 °C for 30 s to denature all double-stranded structures followed by a rapid cooling to 25 °C for 30 s to facilitate successful hybridization between probes and the target strands. The plate was then transferred to a LightScanner (Idaho Technology Inc.). Melting profiles of each well were collected by monitoring the continuous loss of fluorescence with a steady increase of the temperature from 35 to 97 °C with a ramp rate of 0.1 °C/s. The relative quantification was based on the negative first derivative plots using software MATLAB. The areas of probe-target duplexes melting peaks were auto-calculated by ʻAutoFit Peaks I Residualsʼ function in software PeakFit (SeaSolve Software Inc.). The mutant frequency X was calculated using the formula shown below:$$X=frac{{rm{Area}}_{{mathrm{mutant}}}}{{{{mathrm{{Area}}}}}_{{mathrm{{mutant}}}}+{{rm{Area}}}_{{mathrm{{WT}}}}}$$
(1)
To validate the RQ-HRM method, standard curves were generated by measuring mixed samples with known proportions of mutant templates: 0, 10, 20, 30, 40, 50, 60, 70, 80, 90 and 100%. Measurements for each sample were done in triplicate. Linear regression formula of each mutant between actual frequencies and measured frequencies are shown in Fig. S7. The high R2 values, and nearly equal to 1 slope values of these equations, confirmed that the RQ-HRM method can accurately detect mutantsʼ frequency in a mixed population.The relative fitness of the evolved strains was calculated according to previous studies using the following equation75,76:$${mathrm{{relative}}; {mathrm{{fitness}}}}(r)=frac{{{{X}}}_{1}(1-{{{X}}}_{0})}{{{{X}}}_{0}(1-{{{X}}}_{1})}$$
(2)
where X0 is the initial mutant frequency; (1−X0) the initial ancestor frequency; X1 the final mutant frequency; and (1−X1) is the final ancestor frequency.Reporting summaryFurther information on research design is available in the Nature Research Reporting Summary linked to this article. More

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