ABSTRACT Domestication – the artificial selection of wild species to obtain variants with traits of human interest – was integral to the rise of complex societies. The oversupply of food was

probably associated with the formalization of food preservation strategies through microbial fermentation. While considerable literature exists on the antiquity of fermented food, only few

eukaryotic microbes have been studied so far for signs of domestication, less is known for bacteria. Here, we tested if cheese starter cultures harbour typical hallmarks of domestication by

characterising over 100 community samples and over 100 individual strains isolated from historical and modern traditional Swiss cheese starter cultures. We find that cheese starter cultures

reduction of their niche breadth. Future work documenting the prevalence of these hallmarks across diverse fermented food systems and geographic regions will be key to unveiling the joint

history of humanity with fermented food microbes. SIMILAR CONTENT BEING VIEWED BY OTHERS FUNCTIONAL STRAIN REDUNDANCY AND PERSISTENT PHAGE INFECTION IN SWISS HARD CHEESE STARTER CULTURES

Article Open access 06 August 2021 META-ANALYSIS OF CHEESE MICROBIOMES HIGHLIGHTS CONTRIBUTIONS TO MULTIPLE ASPECTS OF QUALITY Article 13 August 2020 LARGE-SCALE GENOME-WIDE ANALYSIS LINKS

LACTIC ACID BACTERIA FROM FOOD WITH THE GUT MICROBIOME Article Open access 25 May 2020 INTRODUCTION Domestication is the process of modifying wild species through artificial selection to the

benefit of a “domesticator”, which is usually human1,2. This process was integral to the rise of complex human societies3,4,5. In particular, the domestication of crop plants6 and

livestock7 over the past 12,000 years4 enabled the emergence of social complexity and initiated large-scale anthropogenic changes in the earth’s biosphere8. Plant and animal domestication is

characterised by phenotypic and genetic changes from the wild ancestor, such as adaptation of body mass9 or change in nutrient content10, which overall contributed to the production of food

surplus. This oversupply of food was associated with the formalisation of food preservation strategies by microbial fermentation – a metabolic process that converts sugars into acids – to

produce, for example, fermented vegetables, wine or cheese11,12. This leads to a decrease in pH, which reduces undesired microbial growth and prevents spoilage of stored food. These

processes were likely adopted by humans many millennia ago13. For cheese, the earliest archaeological evidence can be found within the Neolithic (ca. 12th-5th millennium B.C.)11,13 and

likely became more common and important during the Copper, Bronze and Iron Age (ca. 5th-1st millennium B.C.)14. Fermented food products have diversified in a myriad of forms all over the

globe15,16, constituting healthy and tasty components of the human diet, which are key in many cultures and sustainable opportunities for the future of food in others17. This raises the

fundamental question of whether, as for plants and animals, fermented food microbes have also been domesticated by humans, and if so, when and how this has happened18,19. It is commonly

accepted that some fermented food microbes have been maintained through continuous passaging20,21 and artificial selection for specific traits (e.g., shelf life or taste). Accordingly, we

expect them to present typical “hallmarks” of domestication, i.e., genomic and phenotypic signatures associated with (microbial) domestication that distinguish them from their wild

counterparts. In fact, a handful of microbial domestication cases have been documented empirically22,23, mostly for eukaryotic microbes. For example, _Saccharomyces cerevisiae_ used for

bread making24 and alcoholic fermentation25,26, _Aspergillus oryzae_ used for sake, soy sauce, and miso production27 and _Penicillium camemberti_ found on camembert cheese rind28 show

genomic and phenotypic characteristics that distinguish them from their wild counterparts18,29,30. Thus, while the genomic signatures of domestication are well-defined for eukaryotic

microbes, we know less about the collective genomic and phenotypic consequences of bacterial domestication used for food fermentation29,31. For example, _Oenococcus oeni_, which is

responsible for the malolactic conversion in winemaking, is thought to have rapidly diverged from its ancestor due to the emergence of hypermutator strains, but it remains unclear to what

extent subsequent evolution was influenced by human domestication32. Similarly, strains of _Lacticaseibacillus paracasei_ isolated from ripening cheese show signs of adaptation to milk but

lack other hallmarks of domestication33. Altogether, this suggests that most suspected cases of domestication in fermented food bacteria lack a complete characterisation of domestication

hallmarks previously observed in microbial eukaryotes like yeast. To explain this discrepancy, we posit that, unlike yeasts (i.e., in beer, bread and wine), the previously studied bacteria

have not undergone continuous passaging with rapid and iterative bursts of growth solely on the fermented foods but rather persisted in food-associated environments (e.g., grape skin for

_Oenococcus oeni_). This reduces the strength with which artificial selection can act on evolution. In contrast, cheese starter cultures (starter cultures) are a promising candidate to test

for domestication in fermentation. These cultures have been passaged and selected for thousands of years via backslopping (i.e., continuous re-inoculation of previous-day whey) and have been

extensively selected for flavour and rapid acidification purposes34. Starter cultures are categorised by their usage temperature: mesophilic (below 42 °C) and thermophilic (above 42 °C).

Mesophilic cultures, dominated by _Lactococcus lactis_ or _L. cremoris_, show taxonomic and genetic adaptations to the dairy environment, namely the loss of genes encoding functions needed

in animal and plant environments29. However, these species maintain comparatively large genomes30 and occur in multiple environments31 partially due to the presence of diverse plasmids32,33.

Thermophilic starter cultures are dominated by three thermophilic bacteria, namely _Streptococcus thermophilus_ and _Lactobacillus delbrueckii_ subsp. _lactis_ (hereafter only _L.

delbrueckii_) and _Lactobacillus helveticus_35. Although previous genomic analyses have shown signs of genome decay in these species22,36,37,38,39,40, systematic screening for domestication

hallmarks in these microbial communities is still lacking20,21. In this study, we aimed to detect signs of domestication in thermophilic starter cultures and to date the potential

domestication events. To this end, we collected both modern and historic (1970s) samples of 11 undefined starter cultures that are continuously passaged as undefined starter cultures

(described in refs. 41,42,43,44) and used to make three cheese varieties in different regions in Switzerland (Fig. 1A). We characterised over 1000 samples phenotypically, and about 100

metagenomes and more than 100 bacterial isolates from historical and modern Swiss starter cultures were genetically characterised (Fig. 1B). By conducting species dating and genomic analysis

of additional isolates from public databases, sampled in different geographic locations and dairy fermentations, we expanded on previously proposed hallmarks of domestication for eukaryotes

to define five specific hallmarks of microbial community domestication for bacteria29: (i) phenotypic reliability over time, as a result of the selection for food preservation, (ii) simple

and stable microbial diversity both at the species and strain-level, as a result of continuous passaging, (iii) evolutionary origin of focal species coinciding with the start of food

preservation, (iv) gradual genome decay and (v) adaptation to the food environment by the gradual reduction of niche breadth. Collectively, our results suggest that thermophilic starter

cultures have been domesticated by humans for millennia. RESULTS FOOD PRESERVATION BY RAPID AND RELIABLE ACIDIFICATION AS A RESULT OF LACTOSE FERMENTATION BY THERMOPHILIC CHEESE STARTER

CULTURES The first hallmark of domestication predicts phenotypic stability over time as a result of continuous passaging in a stable environment (i.e., milk) and selection for a specific

trait, especially food preservation but also ripening and flavouring, as determined by a characteristic aroma and favourable process parameters. The underlying phenotypic properties that

give rise to food preservation (i.e., preventing the growth of undesired microbes) are the rapid and reliable (i) acidification of the environment and (ii) reduction of the amount of easily

accessible nutrients (Fig. 1C). We analysed over 1000 weekly routine quality-control measurements for three key phenotypes of food preservation via lactose fermentation, namely

acidification, lactate production, and D- to L-lactate ratio (Fig. 1B, C). In routine starter culture quality control, acidification values above 110 acid base titration value (°Th), amount

of detected lactate between 6 and 12 g/kg and D- to L-lactate ratio between 12 and 45 are generally regarded as efficient milk fermentation. Firstly, we titrated 10 ml of > 1000 samples

and found that acidification potential was consistently (> 110 acid base titration value (°Th)) and reproducibly high over time with no significant temporal trend (Fig. 1D, mixed effect

linear model time slope estimates with _p_ < .05). Secondly, the amount of detected lactate and the ratio of the corresponding enantiomers, D- and L-lactate, slightly but not dramatically

increased over time (Fig. 1E, F, mixed effect linear model time slope estimates with _p_ < .05). The former suggests a stable nutrient depletion by accumulation of lactate as a final

product. The latter suggests that the relative metabolic contribution of the two dominant community members is similar across time. This is essential as _S. thermophilus_ ferments rapidly

but only until pH 5, whereas _L. delbrueckii_ commonly starts significantly fermenting from pH 5 downwards for a longer time. In summary, these results show that starter cultures are

phenotypically stable, measured by rapidly (within the quality control boundaries) and reliably (no significant trend over time, and the distribution of phenotypes within the range of what

is considered efficient in the industry, in Fig. 1d) acidify milk, as expected when communities are selected for specific traits. CHEESE STARTER CULTURES ARE SIMPLE AND STABLE MICROBIAL

COMMUNITIES The second hallmark of domestication predicts that the microbial diversity, both at the species and strain level are simple and stable as a result of the passaging in a contained

and highly stable nutrient-rich environment. We tested this by selecting a total of 98 starter cultures for shotgun sequencing (6–11 Mio. reads per sample; see circles, triangles, and

squares in Fig. 1B) and determined their taxonomic composition using a short read taxonomic profiler (mOTU245). As expected, we found that most samples were dominated by only two species,

_Streptococcus thermophilus_ and _Lactobacillus delbrueckii_ subsp. _lactis_ (Fig. 2A). Yet, we noted two apparent signs of instability over time: (i) _L. helveticus_ was only present in

early samples of some cheese starters, and (ii) the relative abundance of _S. thermophilus_ and _L. delbrueckii_ varied across samples of the same starter culture (Fig. 2A). The former

observation suggests that _L. helveticus_ may have been lost over time without changing the phenotypic properties of the starter cultures. The latter observation may be due to the fact that

the precise sampling time points during the acidification process by the starter cultures were not controlled for and that _L. delbrueckii_ growth is delayed relative to _S. thermophilus_

during acidification41 by their mutualistic interaction termed protocooperation46,47. Also, we cannot exclude that sample degradation through time may have biased the community composition

(the older samples are typically not viable anymore). The stable D- to L-lactate ratio (Fig. 1F) corroborates that the integrated metabolic activity of the two species at the sampling

timepoint remains stable over time. Altogether, this suggests that the species-level composition of the starter culture was remarkably stable over nearly 50 years of sampling, with no

additional species identified in the metagenomic sequences. To assess the within-species diversity, we mapped the metagenomic reads against a reference database containing isolate genomes of

each of the three species and quantified the number of polymorphic sites detected in core genes (i.e., genes identified in all strains of a given species, see methods) in each sample. The

proportion of polymorphic sites was similar over the samples, with around 0.11% (SD = 0.72 %) and 0.02 % (SD = 0.03 %) for _S. thermophilus_ and _L. delbrueckii_, respectively (SFig. 1).

This is comparatively low with respect to bacteria found in non-food fermentation systems like in the gut of animals (3% and 2–10% polymorphic sites within species in the gut microbiota of

human48 and honey bees49, respectively, even for _Lactobacillus_ species). To determine the actual number of strains the detected within-species diversity corresponds to, we genotyped > 

2000 colonies from the 11 starter cultures (Fig. 1B) and sequenced the genomes of 112 isolates. Using an all-against-all genomic distance analysis implemented in poppunk50, we found that the

sequenced genomes separate into 12 _S. thermophilus_, two _L. delbrueckii_ and two _L. helveticus_ sub-species clades (see “Methods”). Overall, the sub-species clades within the different

species are very similar (min ANI: Sterm = 98.6 %, Ldel = 98.9 %, Supplementary Fig. 251). These sub-species clades accounted for most of the SNPs detected by metagenomic sequencing (93% and

78% of the metagenomic SNPs from _S. thermophilus_ and _L. delbrueckii_, respectively, Fig. 2B and Supplementary Fig. 3), suggesting that we have isolated and sequenced most of the

sub-species diversity present in the analysed starter cultures. In the case of _L. delbrueckii_ and _L. helveticus_, a single sub-species clade dominated in all analysed samples, while for

_S. thermophilus_, one to four sub-species clades per sample were detected. Finally, to test if the milk environment in which these bacteria have been passaged cannot carry a larger amount

of diversity at the sub-species level, we inoculated sterile milk with a diverse synthetic community containing all 11 dominant _L. delbrueckii_ isolates (all from the same sub-species

clade) and all 11 dominant _S. thermophilus_ isolates (from seven different sub-species clades) each from one of the 11 starter cultures. We inoculated them at equal amounts and co-cultured

them in five replicates for 27 passages (~ 123 generations) (Fig. 2C). After 21 generations, we found that five of the 11 original _S. thermophilus_ strains were still present. However,

after 123 generations, only one strain could be detected in the community (Fig. 2C), the most dominant strain in RMK 101 and 150). This experiment suggests that only a limited number of

strains can co-exist due to passaging, as observed in natural settings, but that other local factors (not mimicked in our experiment) might contribute to the dominance of alternative strains

in different starter cultures. In summary, these results show that undefined starter cultures have a simple community composition at the species and strain levels that shows a high degree

of stability, in particular at the species level, likely due to the continuous passaging in a stable, closed and nutrient-rich environment. THE ONSET OF CHEESE STARTER STRAIN DIVERSIFICATION

COINCIDES WITH THE ORIGIN OF DAIRY FERMENTATION IN HUMANS The third hallmark of domestication predicts that the evolutionary origin of starter bacteria should coincide with or postdate the

start and diversification of cheese making or dairy fermentation in prehistoric times, as referenced by the archaeological record. The actual timing of the origin of cheese making is

difficult to estimate from archaeological evidence because cheese making tools (wooden vessels, cloth strainers, leather sacks) and residues are usually organic and unlikely to be well

preserved over millennia52,53. The first archaeological evidence of cheese making based on organic matter found in ceramic stainers places the latest estimate of cheese-making origin around

7000 BC in the Near East13 and around 6000 BC in Europe11. We thus hypothesise that the origin of the starter bacteria should similarly fall within the Neolithic, and possibly even into the

Copper, Bronze, and Iron Ages, when important innovations in cheese making occurred11. To test this, we sought to date the evolutionary transition(s) from a non-dairy environment to a dairy

environment using phylogenetic comparative methods. This approach first maps strain or species habitat preference (dairy/non-dairy) onto a phylogeny to identify the evolutionary

transition(s) between habitats and then, in a second step, uses the molecular clock and the dates of historical samples as calibration points to estimate the age of this transition. We

assembled a genomic database of 234 strains from the three cheese starter bacteria as well as closely related species by combining our own dataset with publicly available data (complete list

of genomes provided in Supplementary Data 1). We mapped the isolation source on whole genome phylogenies (for within-species phylogenetic relationships) and on rRNA phylogenies (for

between-species relationships). Most strains from _S. thermophilus, L. delbrueckii_ and _L. helveticus_ were isolated from cheese starter cultures or other milk products, with a few isolates

from faecal samples (Supplementary Fig. 4). Corroborating previous findings54, ancestral niche reconstruction suggests that with a high likelihood of 99.99%, 99.99% and 76% the niche of the

ancestors of all known strains of _S. thermophilus, L. delbrueckii_ subsp. _lactis_, and _L. helveticus_, respectively, were already associated with dairy products (Maximum likelihood

model, Supplementary Fig. 5). In addition, although the three closest related sister species to our focal species are of animal rather than dairy origin, they all encode enzymes to degrade

milk (LacG or LacZ galactosidases, Fig. 3A–C). This suggests that these milk-adapted sister clades probably evolved with the appearance of milk in mammals roughly 200 million years ago55,

but only our focal species are found in the dairy environment. Therefore, we propose that the origin of cheese starter bacteria is located somewhere between the split from the sister species

(stem age) and the most recent common ancestor (MRCA) of the strains within the species (crown age) of the three starter culture species (branches highlighted in black in Fig. 3A–C). To

estimate the age of the non-dairy-to-dairy transition, we took advantage of two independent molecular clocks of the stem and crown age. Stem ages were determined using the divergence time

from the rRNA phylogeny. Assuming an rRNA substitution rate of 1 substitution per site per 100 million years56, we estimated the divergence of _S. thermophilus_, _L. delbrueckii_ subsp_.

lactis_ and _L. helveticus_ from their sister taxon at around 432,000, 264,000 and 1,044,000 years ago, respectively (Fig. 3A–C). While this is more recent than most other species within the

two genera (Fig. 3G), it is still three orders of magnitude older than the first report of dairy fermentation. Crown ages were determined using dated core-genome phylogenies of our focal

species (Fig. 3D–F and Supplementary Fig. 6). Using a molecular clock based on the core genome phylogeny and historical samples as calibration points57, we estimated a substitution rate of

1.1 SNPs per clonal core genome per year, which falls in the average range typically observed for other bacterial species58,59. We extrapolate that the crown age of _S. thermophilus_, _L.

delbrueckii_ subsp. _lactis_ and _L. helveticus_ were around 3221, 7798 and 4304 years ago, respectively (Fig. 3G). This method provides an approximate age range for the different clades

rather than precise dating represented by the large estimated confidence interval errors, ranging from 2163 to 12,708 years ago. Nevertheless, it indicates that the origin of known

strain-level diversity within species is roughly similar to the expected origin of dairy fermentation from archaeological records11. CHEESE STARTER BACTERIA SHOW GENOME DECAY BY TRANSPOSON

EXPANSION The fourth hallmark of domestication predicts that cheese starter bacteria show signs of gene loss and genome decay60. This is expected when bacteria thrive in stable environments

with extensive nutrient availability and relatively small population sizes61. This hallmark additionally predicts that the onset of genome decay must have started after the onset of

domestication itself. To test this hallmark, we compared the genome size and the extent of genome decay (number of pseudogenes) of our focal cheese starter strains to their closest relative.

To put these estimates into a broader context, we also included data on insect-associated bacterial endosymbionts that show extensive genome decay due to pseudogenization62. All three

starter culture species had smaller genome sizes (Wilcoxon test, _p_ < 0.05, Fig. 4A and Supplementary Fig. 7) and a higher number of pseudogenes (Wilcoxon, _p_ < 0.05, Fig. 4A and

Supplementary Fig. 8) than  closely related species. We found that many of the detected pseudogenes in the three starter culture species were the result of insertion events of transposons

(mean = 45%, std = 9%; Supplementary Fig. 9) belonging to 15 different IS element families (Supplementary Fig. 10). Three other _Lactobacillus_ species had similar signs of genome decay, all

of which were also food fermentation-associated species, namely _L. delbrueckii_ subsp. _bulgaricus_, _L. kefiranofaciens_ and _L. amylolyticus_. The observed genome decay of our focal

genomes is in the range of host-restricted symbionts such as _Rickettsia_ spp. and _Wolbachia spp_., but not as extreme as for obligate endosymbionts of plant-sap feeding insects that have

co-evolved with their host for millions of years. Overall, our findings suggest that all three starter culture species have experienced pseudogenization via transposon insertion during their

evolutionary history. If genome decay started at the onset of domestication as predicted from the hallmark, the timing of pseudogenization events should be more recent than the onset of

domestication itself. To estimate the onset of genome decay, we reconstructed the evolutionary history of pseudogenization events by mapping the presence/absence of modern pseudogenes onto

the strain phylogenies. For each pseudogene, we identified the most recent common ancestor (MRCA) of the genomes which contained the pseudogene or in which the gene was completely missing

(assuming the pseudogene was lost, Supplementary Fig. 11). While we observed a few pseudogenes originating around the crown age (oldest peak, Fig. 4B), we found that most pseudogenes likely

originated in the last millennia (intermediate peak, Fig. 4B) or even as recent as in the last century (most recent peak, Fig. 4B). The lag between the crown age (prehistoric) and the

increase in pseudogenization in the last millennia, could arise if cheese making remained a spontaneous fermentation-like process for millennia before being more tightly controlled by

backslopping. We conclude that the overall age similarity between the loss of functional genes (i.e., pseudogenization) and the cheese making development (within the last couple of

millennia) for all three focal species suggests that the persistent bottleneck and selection pressures of cheese-making are associated with recent and ongoing genome decay. REDUCTION OF THE

NICHE BREADTH AND ADAPTATION TO THE CHEESE MAKING ENVIRONMENT The fifth hallmark of domestication predicts a reduction of the niche breadth associated with the loss of non-essential

functions due to the adaptation to the stable and nutrient-rich cheese environment. We first tested the ability of starter culture bacteria to grow on a wide range of carbon sources that are

representative of diverse non-dairy environments. We found that _S. thermophilus_ and _L. delbrueckii_ can metabolise only 5 and 9, respectively, of the 92 tested carbon sources, while

their closest non-dairy relatives could metabolise 12 (58% drop), 61 (85% drop), respectively, (Supplementary Figs. 12, 13). We then sought to explore whether the pseudogenization of genes

observed in these species could explain the loss of these metabolic capabilities. We tested whether cellular functions - in particular carbon metabolism - were enriched in pseudogenes.

Pseudogenes were spread across many orthologous gene families (OGs). Of a total of 19,728 OGs in all three species, we identified 5639 (29%) containing at least one pseudogene. Three COG

categories were overrepresented among the OGs containing pseudogenes, independently in each of the three starter culture species: carbohydrate (G), amino acid (E) and inorganic ion transport

and metabolism (I) (χ2 > 0.3, _p_-value < 0.05, Fig. 5, Supplementary Fig. 14). More specifically, the three KEGG metabolic modules (i) pentose phosphate pathway (ko00030), (ii)

fructose and mannose metabolism (ko00051) and (iii) starch and sucrose metabolism (ko00500) were commonly pseudogenized (Supplementary Fig. 15). To confirm that the identified pseudogenes

are indeed non-functional, we carried out RNA-seq of cheese during the first 24 h of cheese making to look at their expression levels. We found that pseudogenized genes were generally less

expressed than non-pseudogenized genes throughout the first 24 hours of fermentation (RNA-seq experiment, Supplementary Fig. 17). In summary, these results show that recent pseudogenization

affected genes involved in the degradation of carbohydrates that occur in plants but not in milk, suggesting that cheese making strains have lost genes that appear non-essential today in a

stable and predictable dairy environment but were likely more important in their ancestral niche. DISCUSSION Overall, fermented foods represent a stable, microbe-rich environment where

human-controlled continuous passaging and selection results in the establishment of defined and stable microbial communities with specific phenotypic properties. So far, apart from the

domestication of some eukaryotic microbes (e.g., Saccharomyces cerevisiae used in beer, wine or bread), we know surprisingly little about the history of fermented food microbes, and whether

they have been domesticated as extensively as cattle and crop plants remains an open question29,31. We can conclude from our study that thermophilic starter cultures show clear signs of

domestication. (i) They are highly reliable in their acidification and lactose utilisation, likely because of the ongoing selection of the preservation properties (hallmark 1). (ii) They

contain simple and stable microbial communities perhaps because of continuous passaging (hallmark 2). (iii) The timing of the origin of the strains is roughly similar to the emergence of

dairy fermentation and the resulting anthropogenic selection pressure (hallmark 3). (iv) They show clear signs of recent and ongoing genome decay that can be expected from stable and

nutrient-rich environments with relatively small population sizes and continuous passaging (hallmark 4). (v) They show a reduction of the niche breadth and adaptation to the cheese making

environment, suggesting that their current niche is restricted to dairy fermentation batches (hallmark 5). We acknowledge that the domestication hallmarks highlighted here could vary

substantially across species and systems depending on the processes and routes of domestication63. We suggest that comparing and classifying various fermented food systems into distinct

routes of domestication represents an exciting avenue for future research (as previously considered for eukaryotes64). In addition, the list of hallmarks we provide here is not necessarily

complete. For example, recent studies have looked at the evolution of the pangenome of the different clades of _L. delbrueckii_40,65 and _S. thermophilus_39. While these studies similarly

conclude that these species have a distinct evolutionary history tied to dairy fermentation, it currently remains unclear if the pangenome is open and expanding39. Notably, the accessory

genome contains numerous carbohydrate utilisation genes suggesting a broad nutrient range. Here, we suggest that the size of the pangenome is likely also a consequence of extensive and

ongoing pseudogenization because of a substantial fraction of highly mobile genes in the accessory genome, namely active transposases, phages or phage defence genes40,41,66,67.

Interestingly, phages targeting _S. thermophilus_ have adapted to the specific dairy conditions similarly to their host, potentially playing an important role in shaping the remaining

genetic diversity of the bacterial species68. In any case, the addition of more high-quality genomes and a broader sampling in dairy and non-dairy environments will enable us to understand

the pangenome diversity better and explore how it relates to domestication history. Finally, including a wider diversity of cheese starter bacteria from all over the world will probably help

push back the date of the onset of the diversification of cheese bacteria and represent an important research avenue. Collectively, we suggest that thermophilic starter cultures have been

domesticated by humans for millennia. Domestication probably started during the Neolithic era when cheese making emerged but likely gradually continued for millennia14. In our data, this is

evidenced by the large difference in timing of origin between cheese starter members as well as the delay between the origin of the current strain diversity and the pseudogenization. While

the exact timing of the transition from a spontaneous dairy fermentation to backslopping is impossible to pinpoint precisely, our estimates of starter culture origin using molecular data

roughly agree with the archaeological record. Overall, we suggest that the bacteria have likely been in repeated cycles of selection for rapid and reliable acidification and also genetic

drift through the repeat subsampling by backslopping. Our study fills an important knowledge gap by addressing if, when and how microbes have evolved to anthropogenic usage and provide a

conceptual framework that can be applied to other fermented food products. For example, from the pseudogenization analysis (Fig. 4), we pinpoint candidate species that might have been

domesticated, in particular, _L. delbrueckii_ subsp. _bulgaricus_ which has previously been associated with a distinct evolutionary history69 and genomic repertoire40 tightly linked to

yogurt production. Moreover, it will be instrumental in guiding decision makers in the cheese community to decide on optimal community and genomic diversity guidelines, as, for example, has

already been done in beer brewing70. While rapid and reliable acidification is the primary role of starter cultures, many secondary roles, most notably phage defence and flavour production,

play key roles in the selection of starter cultures today. Key questions in domestication research ask whether, for a given domesticate, there was a single domestication event that was

restricted to a particular geographic area that spread across regions or if there were multiple independent domestication events12. For cheese starter bacteria, this remains unknown. In the

case of crop plants and animals, the most likely scenario seems to depend on the domesticate and the continent1,64. The origin of _S. cerevisiae_ is likely Asian, but it is still unclear

whether the domestication happened initially in Asia and the domesticate then spread or whether the wild ancestor spread first and then got domesticated several times in different

places24,71, as reviewed in ref. 29. In the case of cheese starter bacteria, data from across a larger diversity of continents and traditional cheese varieties will be key to differentiating

between a single or a multiple-origin scenario and will probably help push back the date of the onset of diversification of cheese bacteria. In addition, the data we present here do not

encompass closely related “wild” strains that might share a very recent common ancestor with starter culture strains. Discovering and characterising these wild relatives will be instrumental

to reconstructing the evolutionary history of starter culture and testing the single vs. multiple origin scenarios as well as the degree of hybridisation between domesticated and wild

strains. Previous studies have already suggested that these species likely do not have an original niche in the human gut54,72,73,74 potentially having a plant origin. However, we speculate

here that the ancestral niche of cheese starter bacteria might be the gut of milk-feeding (i.e., juvenile) dairy animals that were originally used as a source for rennet75. METHODS COMMUNITY

SAMPLING The starter cultures were continuously passaged at Agroscope (Liebefeld, Switzerland) as undefined starter cultures. The process of maintaining an undefined starter culture is

explained extensively in ref. 41. In short, a stock culture is aliquoted into several samples, which are used on a weekly basis to fulfil the demand of cheese makers. The initial stock is

maintained as a freeze-dried ampule at 4 °C and is only passaged and aliquoted if necessary. The number of passages is irregular and not documented. In general, the stock cultures were

passaged more frequently from 1970 to approximately 2000, as frequently as every month since we have numerous samples dated one month apart. Today, the frequency of passaging is reduced to a

minimum, which can be as rare as every 5 years. The historic samples were stored at 4 °C throughout the years, and DNA isolation was done collectively in 2019. The DNA isolation was done as

previously described76. In short, the DNA was isolated with the EZ1 DNA Tissue kit on the BioRobot EZ1 robot (Qiagen, Hombrechtikon, Switzerland). STRAIN ISOLATION AND BACTERIAL COUNTS

Strains were isolated in 1975 and 2019 (Fig. 1B). The isolates from 1975 were picked from MR11 plates (for both species) and stored in the strain collection of the Agroscope. The isolates

from 2019 were isolated, as previously explained in Somerville et al. 202241. In short, all 11 starter cultures were plated on two selective media SPY9.377 and MR11 (MRS adjusted to pH 5.4

according to ISO7889) for _S. thermophilus_ and _L. delbrueckii_, respectively. Ninety-six colonies per species were randomly picked and cultured in liquid media for 24 h at 37 °C. For

genotyping, DNA from 100 μL of culture was extracted using the EtNa DNA isolation method78, and mini-satellite PCR for strain identification of _S. thermophilus_ and _L. delbrueckii_ was

done as explained previously41. In short, the length of a mini-satellite region in the two species was evaluated by quantifying the PCR length (all primers in Supplementary Data 2) on a

Fragment Analyser™ (Advanced Analytical Technologies, Ankeny, IA, USA). Moreover, colony-forming units (CFU/ml) were determined by serial dilution and plate counting with an Eddy Jet Spiral

Plater and SphereFlash Automatic Colony Counter (both from IUL, Barcelona, Spain). A full description, including all primers used, can be found in the supplemental methods. METAGENOME AND

GENOME SAMPLE PREPARATION AND SEQUENCING Ninety-eight samples, including historic freeze-dried ampules, present working stocks and starter cultures, were prepared for shotgun metagenome

sequencing. The DNA was isolated as previously explained41, and Nextera flex libraries were prepared and subjected to HiSeq4000 150PE (Illumina) sequencing at the Genomic Technologies

Facility in Lausanne, Switzerland. Further, a subset of genomic samples was sequenced on a minION (Nanopore) with a rapid barcoding kit. RAW READ ANALYSIS The raw reads for both metagenomic

and genomic samples were handled similarly. All adaptors and barcodes were removed with trim galore (v0.6.10)79. Reads mapping to the cow genome from the milk was removed with KneadData

(v0.7.3) (https://bitbucket.org/biobakery/kneaddata). The reads were mapped with bwa mem (bwa-0.7.18-r1243)80. The genomes and metagenomes were assembled with SPAdes (v3.13.1)81 for short

reads and Flye for long reads (v2.9.3-b1797)82. Extensive genome polishing was done with Racon (v1.3.3)83 and freebayes (v1.3.7)84. GENOME ANALYSIS The genome assemblies were submitted to

NCBI and annotated with PGAP85. In addition, eggnog mapper was used to identify annotations of pseudogenes86. The pairwise ANI values were calculated with fastANI (v1.33)87. Additional

genomes and their respective PGAP of the three focal species were downloaded from NCBI RefSeq (12.01.2020). Only completely assembled genomes were used as we have previously observed a

substantial number of genes (and pseudogenes) not being assembled with Illumina only assemblies due to the repetitive nature of the tranposase-rich genomes67. Orthofinder (v2.3.1) was used

to identify single-copy core genes within the species88. METAGENOME ANALYSIS The metagenomic raw reads were profiled for species abundance with mOTU2 (v0.3.2)45. In addition, the metagenomic

reads were mapped against reference genomes of the three focal species with bwa mem (bwa-0.7.18-r1243)89. We performed a single nucleotide analysis (SNV) with freebayes-parallel (v1.3.7) on

the genomes and metagenomes in comparison to random reference genomes84. The observed SNVs were filtered with vcftools (v0.1.16)90 and SNPeffect (v4.3t)91 to include only SNVs with a

minimum allele frequency of 0.05, read coverage of 5 and in single-copy-genes (identified with OrthoFinder). The metagenomic SNVs were compared to the previously identified SNVs from the

reference genomes. Sub-species clade frequency was calculated by averaging the allele frequency of all sub-species clade-specific SNVs (for details, see script). PROPAGATION EXPERIMENT AND

MEASUREMENTS For the propagation experiment, we created a pooled starting sample consisting of one random isolate of _L. delbrueckii_ and one of _S.thermophilus_ per starter culture (in

total 22 isolates) (see selection in Supplementary Data 1). The starting sample was propagated in five replicates as described in Somerville et al. 202241. In short, the samples were

propagated to simulate the production of starter cultures. We conducted two passages per week. On the first day, 100 µl of the freeze-dried sample was inoculated into 10 ml autoclaved

organic milk media (BM) and incubated for 18 h at 37 °C. For the second passage on the next day, the pre-culture was inoculated into 10 ml autoclaved BM and incubated for another 18 h at 37 

°C. For the final step, 100 µl of the incubated samples were transferred into a freeze-dry ampule and stored at − 30 °C for at least 1 h. Thereafter, the samples were freeze-dried for 7 

hours until dry. For pH measurements, we used the hydroplate system (PreSens, Germany). The pH was normalised with pH standards of pH 4 and 7. The measurements were done in four replicates

for 30 h at 37 °C. SPECIES-LEVEL LACTIC ACID BACTERIA PHYLOGENY All representative genomes from lactic acid bacteria species were downloaded from NCBI RefSeq. Moreover, from the genus

_Streptococcus_ and _Lactobacillus_, all genomes deposited on NCBI before 12.01.2020 were included. The annotations were screened for Galactosidase (lacZ and lacG). The phylogeny was

reconstructed by concatenating the 16S, 23S and 5S rRNA sequence of the genomes, making a multi-sequence-alignment file with mafft-linsi (v7.526)92 and calculating the phylogeny with RaxML

(v8.2.12) and the “GTRCAT” model93. The plot was created in R with ggtree94. STRAIN-LEVEL DATED WHOLE GENOME PHYLOGENY The preliminary species tree was calculated with OrthoFinder

(v2.3.1)88. Therefore, we back-translated the single copy core genes into the nucleotide space and created a core genome species tree and gene trees as described in ref. 95 using MAFFT

(v7.526)96 and RAxML (v8.2.12) with 100 bootstrap rounds and the GTRCAT93. Moreover, by including the gene tree and the species tree we calculated the clonal species tree with ClonalFrameML

(v1.12)97. Therefore a dated phylogeny was calculated with BactDating (v1.1.2) and 107 Bayes repetitions using our historical samples as time calibration points57. From the subsequent

phylogeny, we predicted sub-species clades for our genome isolates with poppunk (v1.2.0)50. In order to quantify the presence of sub-species clades in the metagenomes, we identified all

sub-species clade-specific SNVs in the core genes. The dated phylogeny and the information from the isolation source (Supplementary Data 1) were used to reconstruct ancestral habitat

reconstruction with the ace function from the ape package98. TRANSCRIPTOME ANALYSIS Samples for transcriptome analysis were collected throughout the first 24 hours of a regular gruyere-type

cheese making process at the cheese pilot plant at Agroscope (Liebefeld, Switzerland). The samples were immediately stored in liquid nitrogen and the RNA extraction was carried out with the

Qiagen EZ1 extraction robot and the RNA Tissue kit. Illumina libraries were prepared with the TruSeq Str-RNA Zero, and sequencing was performed on a 150 PE HiSeq 4000 (Illumina) at the

Genomic Technologies Facility in Lausanne, Switzerland. The Illumina sequences were cleaned with trim galore (v0.6.10)79 and sortmeRNA (v2.1)99. Mapping of reads to isolate reference genomes

was performed with bwa mem (bwa-0.7.18-r1243)89 and gene counting with HT-seq (v0.11.2)100. Further, sample and gene normalisation was performed with DESeq2101. PHENOTYPIC ASSAYS OF

ISOLATES The carbohydrate utilisation profiles were determined using Biolog™ phenotypic microarray plates PM01 and PM02A. Samples were prepared according to the manufacturer’s protocols.

Plates were incubated in Omnilog™ for 48 h at 37 °C. The equipment records the contrast difference every 15 min to generate growth curves. This data was evaluated using Biologs data

acquisition software™ and the opm package in ref. 102. PHENOTYPIC ASSAYS OF STARTER CULTURES The starter cultures were regularly monitored as part of the regular in-house quality control at

Agroscope starting in 1996 (acidity, lactate ratio) and 2000 (total lactate) until 2020 (last measurement included in the manuscript). Specifically, (1) the sample acidity was measured

weekly, (2) the total lactate and (3) the lactate ratio was measured monthly. The acidity of the cultures was determined as follows: 10 mL reconstituted skim milk was inoculated using 0.1 ‰

(10 µl) of the corresponding culture and incubated for 18 h at 38 °C. 1 drop of phenolphthalein was added to the sample and then titrated with 0.1 M NaOH till a visible colour change was

detected. The recorded volume of 0.1 M NaOH in mL was multiplied by 10 and rounded to the whole number resulting in the determined °Th (or Clark degree). Total lactate (D- and L-Lactate) was

analysed enzymatically according to the instruction protocol of the kit manufacturer (Boehringer, Manheim, Germany) using an automated spectrophotometric analyser (Gallery, Thermo,

Switzerland). The proportion of L-lactic acid to total lactic acid was calculated as a percentage. This method has previously been published103. The linear mixed effects model was fitted

into the phenotypic data with the lmer function in the lme4 package104. STATISTICS AND REPRODUCIBILITY No statistical methods were used to predetermine sample size, but rather all available

samples were used. The experiments and lab work were, wherever possible, randomised and blinded. If not mentioned otherwise, the data analyses and statistics were done in R (R Core Team,

2020) and plotting was done using ggplot2105. The wilcox.test function from the rstatix package and all other tests (_t_ test and Pearson chi-square) from the base package was used. All

details and specific parameters can be retrieved in the available code (see “Code availability” section). Moreover, the version numbers for all tools used in this study are provided in the

supplemental methods. REPORTING SUMMARY Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article. DATA AVAILABILITY Raw sequence

data and, if applicable, assembled genomes that support the findings of this study have been deposited in on NCBI with the BioProject codes: PRJNA717134 (Illumina only genomes), PRJNA717134,

PRJNA1083966 (ONT and Illumina genomes), PRJNA1048529 (shotgun metagenomics) and PRJNA1157897 (RNA-seq). In addition, all data frames, intermediate figures and outputs are available on

Zenodo (https://doi.org/10.5281/zenodo.10783580). CODE AVAILABILITY The complete code is available on Git Hub (https://github.com/Freevini/Starter-Culture-diversity) or as a permanent

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Didelot for the useful advice for calculating the molecular clock, Herwig Bachmann and Bas Teusink for the many discussions on these topics, Julien Marquis and Johann Weber from the Genomic

Technologies Facility in Lausanne, Alban Ramette from the IFIK for the sequencing. The authors would like to thank SNF Doc.Mobility P1LAP3_199476 (V.S.), SNF Postdoc.Mobility P500PB_214419

(V.S.). This work was supported as a part of NCCR Microbiomes, a National Centre of Competence in Research, funded by the Swiss National Science Foundation (grant number 180575). F.M. thanks

Uni. of Lausanne, NCCR Microbiome, SNF and Antoine Guisan. AUTHOR INFORMATION Author notes * These authors contributed equally: Florent Mazel, Philipp Engel. AUTHORS AND AFFILIATIONS *

Department of Fundamental Microbiology, University of Lausanne, Lausanne, Switzerland Vincent Somerville, Florent Mazel & Philipp Engel * Agroscope, Liebefeld, Switzerland Vincent

Somerville, Nadine Thierer, Remo S. Schmidt, Alexandra Roetschi, Lauriane Braillard, Monika Haueter, Hélène Berthoud, Noam Shani & Ueli von Ah * Université Laval, Quebec, Canada Vincent

Somerville * McGill, Montréal, Canada Vincent Somerville Authors * Vincent Somerville View author publications You can also search for this author inPubMed Google Scholar * Nadine Thierer

View author publications You can also search for this author inPubMed Google Scholar * Remo S. Schmidt View author publications You can also search for this author inPubMed Google Scholar *

Alexandra Roetschi View author publications You can also search for this author inPubMed Google Scholar * Lauriane Braillard View author publications You can also search for this author

inPubMed Google Scholar * Monika Haueter View author publications You can also search for this author inPubMed Google Scholar * Hélène Berthoud View author publications You can also search

for this author inPubMed Google Scholar * Noam Shani View author publications You can also search for this author inPubMed Google Scholar * Ueli von Ah View author publications You can also

search for this author inPubMed Google Scholar * Florent Mazel View author publications You can also search for this author inPubMed Google Scholar * Philipp Engel View author publications

You can also search for this author inPubMed Google Scholar CONTRIBUTIONS V.S.: Conceptualisation, formal analysis, Funding Acquisition, Visualisation, Writing – Review & Editing,

U.v.A., F.M., and P.E.: Funding Acquisition, Writing, Conceptualisation – Review & Editing. N.T., A.R., L.B., M.H., H.B., N.S., and R.S.S.: Methodology. CORRESPONDING AUTHOR

Correspondence to Vincent Somerville. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests. PEER REVIEW PEER REVIEW INFORMATION _Nature Communications_ thanks

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