Rapid evolution of an overt metabolic defect restores balanced growth

In eukaryotes, conserved mechanisms ensure that cell growth is coordinated with nutrient availability. Overactive growth during nutrient limitation leads to rapid cell death. Here, we demonstrate that cells can adapt to this nutrient-growth imbalance by evolving major metabolic defects. Specifically, when yeast lysine auxotrophic mutant lys− suffered nutrient-growth imbalance in limited lysine, a sub-population repeatedly evolved to lose the ability to synthesize organosulfurs (lys−orgS−). Organosulfurs, mainly glutathione and glutathione conjugates, were released by lys− cells during lysine limitation when nutrient-growth is imbalanced, but not during glucose limitation when nutrient-growth is balanced. Limiting organosulfurs conferred a frequency-dependent fitness advantage to lys−orgS− by eliciting a proper slow growth program including autophagy. Thus, nutrient-growth imbalance can trigger rapid niche construction, which in turn enables the selection of an overt metabolic defect to better tune cell growth to the metabolic environment.


Introduction
All organisms must sense the availability of nutrients. In eukaryotes, when nutrients are abundant, cells express growth-promoting genes associated with, for example, protein synthesis and mitochondrial function, and grow 1 . When nutrients are limited, cells launch stress-response programs and undergo, for example, quiescence and autophagy 1,2 . Mechanisms that ensure this coordination between nutrient availability and growth, or nutrient-growth balance, are conserved across eukaryotes 3 .
Slowing growth during nutrient limitation is essential for cell survival. If cells attempt to grow despite being limited for essential metabolites, they could suffer rapid death from this nutrient-growth imbalance. For example in S. cerevisiae, when a nutrient required by all cells (e.g. carbon 4 , nitrogen 5 , or sulfur 6 ) is limiting ("natural limitation"), cells slow down growth and thus survive. In contrast, when an auxotrophic mutant is limited for the essential metabolite that it has lost the ability to make ("unnatural limitation"), often the mutant fails to slow down growth and consequently suffers significantly reduced viability compared to viability during natural limitation 7,8 . Nutrient-growth imbalance is in part due to erroneous activation of the growth activator TORC1 (Target of Rapamycin Complex 1) during unnatural limitation. Inactivating TORC1, which can be achieved by loss-of-function mutations in the TORC1 pathway or by applying the TORC1 inhibitor rapamycin, restores cell viability 7 .
Since nutrient-growth balance is regulated in a conserved fashion across eukaryotes, and since some cancers are believed to suffer nutrient-growth imbalance 9 , we investigate how cells suffering nutrient-growth imbalance might evolve, taking advantage of a yeast auxotroph mutant.

Nutrient-growth imbalance in lys-cells during lysine limitation
To examine how cells might cope with nutrient-growth imbalance, we took advantage of a lysine auxotrophic yeast strain where the LYS2 gene has been deleted ("lys -"). Consistent with previous work 8 , lys cells starved for lysine died rapidly (Fig 1A, dark blue circles), although lys cells simultaneously starved for lysine and carbon died slowly (Fig 1A, green circles). This is presumably because carbon shortage allowed cells to enter the proper state of no growth (Fig 1A, ii), while lysine starvationan unnatural limitationwas not properly sensed, and consequently cells would attempt to grow despite lysine shortage and die (Fig 1A, iii). Expression of genes associated with autophagy, a process where the degradation and recycling of cellular components facilitates stress survival, was reduced during nutrientgrowth imbalance, which could also contribute to low cell viability 8 . Indeed, poor cell viability was rescued by rapamycin which inhibited TORC1, a growth promoter and autophagy inhibitor ( Fig 1A, blue crosses; i). In summary, lys cells limited for lysine suffered nutrient-growth imbalance and poor viability.

Evolution of organosulfur auxotrophy in nutrient-growth dysregulated cells
To examine how cells might cope with nutrient-growth imbalance, we evolved lys cells in lysine-limited chemostats for tens of generations (Methods, "Evolution"; "Chemostats and turbidostats"). We then randomly chose a total of ~70 evolved clones for whole-genome sequencing. Similar to our previous findings 10 , each clone carried at least one mutation that increased the cell's affinity for lysine. These mutations included gaining an extra copy of chromosome 14 which harbors the high-affinity lysine permease gene LYP1 12 , or losing or reducing activities of genes involved in degrading Lyp1 (e.g. ECM21, RSP5, and DOA4) 11 (Table S2).
Surprisingly, even though the medium originally contained no organosulfurs (sulfur-containing organic molecules; Fig S1), ~10% of the ~70 sequenced clones also harbored mutations in the biosynthetic pathway that converts externally-supplied sulfate to organosulfurs essential to cell viability. These mutations included met10, met14, and met17 ( Fig 1B). Indeed, these mutant clones required an external source of organosulfur such as methionine to grow (Fig 1B), consistent with previous findings 13 . We designate these mutants as lys -orgS -. Since our growth medium was not supplemented with any organosulfurs, lys -orgS cells must have received organosulfur(s) from lys cells in the same culture which are capable of synthesizing organosulfurs. Since lys and lys -orgS clones from the same culture harbored identical ecm21 or rsp5 mutation (Table S2, matched color shading), ecm21 or rsp5 mutation arose first, followed by orgS -. When we tested three independent chemostat cultures and three independent crossfeeding communities at 30~80 generations, lys -orgS mutants could be detected in all cases (Fig S2; Methods "Quantifying auxotroph frequency").

The organosulfur niche mainly consists of glutathione S-conjugates and glutathione
We sought to identify the released organosulfurs ("organosulfur niche") in chemostat supernatants. When ancestral lys cells were cultured in lysine-limited chemostats, organosulfurs rapidly accumulated in the supernatant and reached a steady-state ( Fig S3). Using liquid chromatography-mass spectrometry (Methods, "LC-MS"), we identified glutathione as a released organosulfur (Fig 2A). Glutathione (GSH) is a tri-peptide comprising glutamate, cysteine (containing -SH), and glycine. In the cell, GSH is a major redox buffer, and can form sulfur-conjugates with itself (GS-SG) and with other compounds via disulfide bonds (e.g. GS-S-CoA) 14 . Consistent with LC-MS results, gsh1-cells, whose growth can be supported by externally-supplied GSH or GSH S-conjugates (GS-X) but not methionine (Fig S1, Fig S4) 15 , grew in chemostat supernatants (Fig S4E).
Further chemical analysis revealed that the organosulfur niche contained mainly GS-X and some GSH. Specifically, we estimated the total organosulfur niche using a yield-based bioassay (Methods, "Bioassays"). We used met10 -, a mutant isolated in our evolution experiment which can use GSH, GS-X, and non-GSH organosulfurs such as methionine ( Fig S4). By comparing the final turbidity of met10 in a chemostat supernatant versus in various known amounts of GSH, we can estimate the organosulfur niche in terms of GSH equivalents (Fig 2B, i). We then quantified GSH using HPLC and LC-MS ( Fig 2B, ii and iii; Methods, "HPLC", "LC-MS"). Finally, we quantified the total abundance of GSH and GS-X by chemically reducing GS-X to GSH (Methods) and then measuring total GSH via LC-MS (Fig 2B, iv). GSH and GS-X together dominated the organosulfur niche ( Fig 2B, compare i and iv), with GS-X being the major component ( Fig 2B, compare iii with iv).
The organosulfur niche was mainly created by live cell release rather than by cell lysis. Specifically, we compared the organosulfur concentration quantified in supernatant (Fig 2C, brown) with that attributable to dead cell release (Fig 2C, purple; calculated by multiplying intracellular organosulfur content with dead cell density). Our measurements suggest that organosulfurs were mainly released by live cells ( Fig  2C; Fig S6), consistent with the observation that GSH and GS-X are exported from the cell in an ATPdependent fashion 14,16 .

Organosulfur release is associated with nutrient-growth imbalance
To test whether organosulfur release is associated with nutrient-growth imbalance or with slow growth, we grew lys cells in glucose-limited chemostats and lysine-limited chemostats at the same doubling time.
In contrast to lysine limitation, in glucose limitation nutrient-growth balance is normal and percent dead cells is lower (Fig 1A; Fig 3A). Organosulfurs accumulated to steady state in lysine-limited chemostats but were undetectable in glucose-limited chemostats, despite the latter's higher steady-state live population density (Fig 3). As a further test, we grew lys-cells in nutrient-rich turbidostats 17 (Methods, "Chemostats and turbidostats") where cells grow fast in excess nutrients and thus nutrient-growth balance is achieved. lys2-cells in nutrient-rich turbidostats released GSH at a much slower rate than in lysinelimited chemostats (Fig S5). Taken together, these results suggest that organosulfur release is associated with nutrient-growth imbalance.

Organosulfur limitation confers a frequency-dependent fitness advantage to orgS -
How might lys -orgS rise from the original mutant cell to a detectable frequency during lysine limitation?
To directly test whether orgS mutation conferred fitness benefit to lys-cells, we restored met10 to MET10 in an evolved lys -orgS clone, and compared isogenic lys -orgS and lys clones in a variety of nutrient environments. According to the prevalent "energy saving" hypothesis 18,19 , lys -orgS might be spared of the cost of de novo synthesis of organosulfurs by utilizing external organosulfurs. If this was true, then excess organosulfurs should help lys -orgS to perhaps be even more fit than lys -. However, we observed the opposite: when both lysine and GSH were in excess, lys -orgS grew significantly slower than lys - (Fig 4A). During lysine limitation, lys -orgS survived better than lys during organosulfur limitation  Fig S8). A similar trend was observed at intermediate levels of lysine and GSH ( Fig S9).
An alternative hypothesis is that organosulfur limitation restored nutrient-growth balance in lys -orgS during lysine limitation, thus conferring lys -orgS a fitness advantage over the nutrient-growth imbalanced lys -. That is, organosulfur limitation posed a "natural limitation" which properly shut down growth as well as turned on stress-response pathways such as autophagy in lys -orgS cells. Consistent with this notion, the poor survival of lys -orgS in high organosulfur was rescued by growth inhibitor rapamycin ( Fig 4B orange cross; Fig S8). Deletion of ATG5, a gene essential for autophagy, led to poor survival of lys -orgS cells despite growth inhibition by rapamycin ( Fig 4B green; Fig S8).
To directly test any fitness difference between isogenic lys and lys -orgS clones, we marked them with different fluorescent proteins, and competed them in lysine limitation (Methods. "Competition"). When lys -orgS was initially abundant, its frequency declined as expected (Fig 4C purple) due to competition for the limited organosulfurs released from rare lys -. When lys -orgS was rare, its frequency initially increased (Fig 4 C red and green), demonstrating a fitness advantage over lys -. Regardless of the initial value, the strain ratio converged to the steady state (Fig 4). The observed steady state ratio is consistent with organosulfur release and consumption measurements (Methods,"Competition"). Taken together, by restoring nutrient-growth balance, organosulfur limitation confers lys -orgS a negative frequencydependent fitness advantage over lys during lysine limitation.

Discussions
Our work demonstrates that a cell population with nutrient-growth imbalance (lys cells in lysine limitation; Fig 1A) often diverged into two subpopulations within tens of generations: lys and lys -orgS -( Fig 1B and Fig S2). The probable sequence of events is the following: Upon lysine limitation, cells initially adapted to lysine limitation via mutations such as ecm21, rsp5, and Disomy14 that increased the membrane abundance of the lysine permease Lyp1, and such mutations would become fixed in the culture. Live lys cells released organosulfurs consisting primarily of GS-X and, to a less extent, GSH (Figs 2, S3, S5, and S6). This created an organosulfur niche. When lys -orgS -(e.g. lys -met10 -) arose by mutation, it could grow in the organosulfur niche ( Fig S4). Initially, lys -orgS had a fitness advantage over lys - (Fig 4 and Fig ). This is presumably because for lys -orgS cells, low concentrations of organosulfur mimicked natural sulfur limitation and thus slowed the over-active growth. Indeed, lys -orgS survived lysine limitation better than lys - (Figs 4 and S8), and the superior survival of lys -orgS required autophagy (Figs 5 and S8). As lys -orgS increased to a higher frequency, its fitness advantage over lys would diminish due to the increasing ratio of organosulfur consumer to organosulfur supplier. Eventually, a steady-state ratio of lys -orgS -:lys could be reached (Fig 4), until new mutations alter the relative fitness between the two populations ( Fig S2).
Chemical release by cells ("niche construction") mediates diverse microbial interactions [20][21][22][23] . Indeed, auxotrophs are predicted to be widespread 19,24 and have been found in natural isolates [25][26][27][28][29][30] . Often, metabolites are released as a consequence of "overflow" metabolism [31][32][33][34][35] . For example, yeast excretes amino acids as a consequence of nitrogen overflow, and the released amino acids in turn enable the survival of symbiotic lactic acid bacteria 32 . The release of GSH and GS-X by lysine-limited lys cells is likely a consequence of nutrient-growth imbalance (Fig 3, Fig S5C). GSH is the major cellular redox buffer in eukaryotes 36 . For example under conditions of heat stress, GSH is known to react with proteins to form protein-S-SG disulfides, which protects the cell against damage from the formation of disulfide bridges between proteins 37 . Both GSH and GS-X are known to be actively exported from cells. For example, in higher eukaryotes, GSH is exported from the liver and transported to other organs via the blood stream, and GS-SG is exported for biliary excretion. Our results suggest that GS-X and GSH are likely released by live cells (Fig 2C; Fig S6), consistent with the ATP-dependent export of GSH and GS-X previously observed in yeast 16 . Furthermore, our data suggest that compared to GSH, GS-X might be preferentially exported (Fig 2C), consistent with cells purging unwanted GS-X. Our findings are consistent with previous observations that yeast upregulated glutathione transporters such as Gex1 and excreted GSH and GS-X 38 in response to cadmium-induced redox stress, and that Opt1, an oligopeptide transporter shown to export GSH 39 , was upregulated in response to different environmental stressors 40,41 .
Evolution of auxotrophy has been experimentally observed in previous work. At a high mutation rate (e.g. a high loss rate of plasmids that carry biosynthetic genes), auxotrophs could rise to a detectable level 42 possibly due to mutation-selection balance. When nutrients were supplied by either the medium or other microbes, initially rare auxotrophic mutants could rise to a detectable frequency due to their fitness advantages over their non-auxotrophic counterparts 18,19,43,44 . In the "energy saving" hypothesis, this fitness advantage is thought to be due to auxotrophs saving the energy of synthesizing the essential nutrient 18 . Contrary to the energy-saving hypothesis, the fitness advantage of lys -orgS over lys was abolished when excess GSH was supplemented (Fig 4; Fig S8). Instead, the fitness advantage of orgSwas only present when organosulfurs were present at a low level. Limited organosulfurs, by mimicking natural sulfur limitation and shutting down the inappropriately active growth in lys -orgS -, could allow these cells to survive lysine limitation better than lys - (Fig 4).
In summary, lysine-limited lys cells release organosulfurs. These organosulfurs at low levels not only support the growth of rare lys -orgS mutants, but also restore their growth regulation by imposing natural limitation. Consequently, lys -orgS gain an advantage over lys and rise in frequency up to a steady-state value (Fig 4). Interestingly, nutrient-growth imbalance has been implicated in certain cancers 9 . Thus, understanding the evolutionary and ecological consequences of nutrient-growth imbalance in mammalian cells could be fruitful. convention. For all experiments, frozen yeast strains stored at -80 o C were first struck onto YPD (10 g/L yeast extract, 20 g/L peptone, 20 g/L glucose) + 2% agar plates and grown at 30 o C for ~48 hours, from which a single colony was inoculated into 3 mL of YPD and grown overnight at 30 o C with agitation. All experiments were carried out within 3 days of generating the overnight culture. Minimal medium (SD) contained 6.7 g/L Difco Yeast Nitrogen Base w/o amino acids w/ ammonium sulfate and 20 g/L Dglucose, except during glucose-limitation experiments where lower levels of glucose were used as specified. Depending on strain auxotrophy, SD was supplemented with lysine (164.3 µM), adenine sulfate (108.6 µM) 45 , or organosulfurs (134 µM) so that cells could grow exponentially.

Evolution
Coculture evolution was described in an earlier study 12 . To revive a coculture, ~20 µL was scooped from the frozen sample using a sterile metal spatula, diluted ~10-fold into SD, and allowed to grow to moderate turbidity for 1-2 days. The coculture was further expanded by adding 3 mL of SD. lys-evolved clones were isolated by plating the coculture on rich media (YPD) agar with Hygromycin B.
For monoculture evolution, chemostat vessels ( Fig S10) were placed into a core manifold with six receptacles, each with a magnetic stirrer. Reactor vessels were immobilized in receptacles by adjustable compression rings. A rubber stopper equipped with an inflow tube and a sampling needle covered the top of the vessel. Waste flowed by gravity to a waste receptacle below the device through C-Flex 0.375" ID tubing (Cole Parmer) attached to the outflow arm. Nutrient media was fed to the vessels from media reservoirs by tubing passed through a Cole Parmer MasterFlex C/L peristaltic pump controlled by a custom LabView console through a custom relay box. Media reservoirs were 1 L glass bottles capped with one-hole rubber stoppers. Tubing was fed through the stopper and allowed to hang to the bottom of the reservoir. The reservoirs were placed on Denver Instrument XP-1500 digital balances, and the actual flow rate for each vessel was determined from the rate of mass loss of the corresponding reservoir.
To create a sterile environment, initial assembly was done in autoclave trays, with vessels held in tube racks. Six reservoirs were prepared by adding 810 mL water to each bottle. Six vessels were prepared by adding a 10 mm stir bar and 20 mL growth media (SD+21 µM lysine) to each vessel. Media delivery tubing was attached between reservoirs and vessels through rubber stoppers, and waste tubing was attached to each outflow arm, with the unattached end covered by foil held in place by autoclave tape. A 1.5 mL micro-centrifuge tube was placed over the sampling needle and held in place by autoclave tape. Tubing ports were wrapped with foil as well. Each reservoir with its attached tubing was weighed, the entire assembly autoclaved, then each reservoir weighed again. Lost water was calculated and added back. Under sterile conditions, 90 mL of 10X SD and a lysine stock were added to each reservoir to reach a final lysine concentration of 21 µM. The vessels were then secured into the chemostat manifold receptacles, reservoirs placed on the scales, and tubing threaded into the pumps.
Ancestral or evolved lys-clones were grown in 50 mL SD + 164 µM lysine for ~20 hours prior to inoculation. Before each experiment, growth was tracked to ensure cells were growing optimally (~1.6 hour doubling time). When cells reached a density of ~0.2 OD600, cells were washed 3 times in SD and inoculated in a chemostat vessel prefilled with SD + 21 µM lysine. After this step, chemostat pumps were turned on at a set doubling time in the custom written LabView software package. Each chemostat vessel contained ~43 mL running volume, and was set to a target doubling time (e.g. for 7 hour doubling, flow rate is 43*ln2/7=~4.25 mL/hr). With 21 µM lysine in the reservoir, the target steady-state cell density was 7x10 6 / mL. In reality, live cell densities varied between 4x10 6 /ml and 1.2x10 7 /ml. Periodically, 4 mL of supernatant was harvested and dispensed into a sterile 15 mL conical tube. Next, 300 µL of this cell sample was removed and kept on ice for flow cytometry analysis. The remaining 3.7 mL of supernatant was filtered through a 0.22 µM nylon filter into 500 µL aliquots and frozen at -80C. Each chemostat was sampled according to a pre-set sequence. For experiments with metabolite extraction, the chemostat vessel stopper was removed and cells from 20 mL of sample was harvested. Due to the breaking of vessel sterility, this would mark the end of the chemostat experiment.
The nutrient reservoir was refilled when necessary by injecting media through a sterile 0.2 micron filter through a 60 mL syringe. To take samples sterilely, the covering tube on the sampling needle was carefully lifted, and a sterile 5 mL -syringe was attached to the needle. The needle was then wiped with 95% ethanol, and slowly pushed down so that the tip was at least ~10 mm below the liquid level. A 5 mL sample was drawn into the syringe, the needle pulled up above the liquid surface, and an additional 1 mL of air drawn through to clear the needle of liquid residue. The syringe was then detached and the cap placed back on the needle. The samples were ejected into sterile 13 mm culture tubes for freezing and flow cytometry determination of live cell densities. In both evolution experiments, samples were frozen in 1 part 20% trehalose in 50mM sodium phosphate buffer pH 6.0 + 1 part YPD. The samples were cooled at 4°C for 15 min before frozen down at -80°C.
Whole-genome sequencing and data analysis are described in detail in 12 .

Quantifying auxotroph frequency
Frozen cultures (2 time points from three mono-culture evolution and three co-culture evolution experiments) were revived, and clones were isolated and screened for auxotrophy. We revived frozen samples by directly plating samples on YPD (monoculture) or YPD + hygromycin (cocultures to select against the partner strain). Plates were grown at 30 o C for 2~4 days until all colonies were easily visible for picking. We observed a variety of colony sizes, and screened both large and small sized colonies when both were present. We counted large and small colonies to estimate the ratio of large:small colonyforming cells in the population, then multiplied this fraction by the fraction of auxotrophs observed in each colony size class to get a full population auxotroph frequency estimate. To screen for auxotrophy, entire colonies were inoculated into 150 µl of SD, 10 µl of which was diluted into 150 µl SD in microtiter plates and incubated overnight to deplete organosulfur carry-over or cellular organosulfur storage. In the case of some small colonies, no dilution was made as the inoculated cell density was already low enough, based on OD measurements. 10-30 µl were then diluted into a final volume of 150 µl each of SD+Lys, SD+Lys+Met, and YPD, aiming for OD~0.005-0.03 based off an initial reading by a 96-well plate OD600 reading of the starvation plate. Plates were then incubated for 48+ hours to grow cultures to saturation, and culture turbidity (OD600) was read using a 96-well plate reader. Control wells of known lys -orgS -(WY1604) and lys -(WY2226) were included in the screening as controls. Wells that grew in SD+Lys+Met and YPD but failed to grow in SD+Lys were scored as lys -orgS -.

Fluorescence microscopy
Fluorescence microscopy experiments and data analysis are described in detail elsewhere 46 . Briefly, the microscope is connected to a cooled CCD camera for fluorescence and transmitted light imaging. The microscope harbors a temperature-controlled chamber set to 30 o C. The microscope is equipped with motorized switchable filter cubes capable of detecting a variety of fluorophores. It also has motorized stages to allow z-autofocusing and systematic xy-scanning of locations in microplate wells. Image acquisition is done with an in-house LabVIEW program, incorporating autofocusing in bright field and automatic exposure adjustment during fluorescence imaging to avoid saturation. Previous analysis 46 has demonstrated that background-subtracted fluorescence intensity scales proportionally with live cell density, and a decrease in fluorescence intensity correlates well with cell death.

Chemostats and turbidostats
Cells were grown under controlled conditions using a custom-made continuous culturing device ( Fig  S10A), with six channels (Fig S10C) where each can be independently operated as a chemostat or a turbidostat. When operated as a chemostat, a channel provides a limited nutrient environment where the growth rate is held constant. When operated as a turbidostat, a channel maintains a constant cell density while cells grow with an abundant supply of nutrients.
The continuous culturing device consisted of six reactor vessels (Fig S10A), each with a volume of ~43ml determined by the height of the outflow tube (Fig S10C, E). A rubber stopper equipped with an inflow tube and a sampling needle covered the top of each vessel (Fig S10C). The vessels were placed in an aluminum mounting frame with six receptacles (Fig S10A, back), each equipped with an integrated magnetic stirrer (made from a CPU fan, magnets, and 3D printed clip) and an LED-phototransistor optical detector for OD measurements. The vessels were immobilized in the receptacles by adjustable compression rings. A sampling needle passed through a seal provided by a short length of glass tubing, so that sampling needle could be easily lowered into the reactor for sampling and elevated above the culture level when not sampling. The tightness of the seal was adjusted by application of zip-ties to the end of glass tubing with the proper tightness to allow movement yet with enough friction to maintain position. Waste flowed by gravity to a waste receptacle below the device through 0.375" ID tubing (Cole Parmer C-Flex) attached to the outflow arm. Nutrient media was fed to each vessel from an independent reservoir by a peristaltic pump (Welco WPM1, operated at 7V DC). The media delivery tube consisted of two sections of generic 2mm OD, 1mm ID PTFE tubing, joined by insertion into the two ends of a 17cm section of PharMed AY242409 tubing (Saint-Gobain) which was inserted into the peristaltic pump ( Fig  S10B). The pump was activated and deactivated by the custom LabView program through a relay box (Pencom Design, Inc. UB-RLY-ISO-EXT-LR). Depending on whether a given channel was in chemostat or turbidostat mode, the LabView program controlled pump in different ways (i.e. constant dilution rate in chemostat, or dilution to a set turbidity in turbidostat). Data for OD and flow rate data were logged in either mode of operation. In both cases, flow rate can be used to calculate growth rate. Media reservoirs (Fig S10A front) were 1 L glass bottles capped with one-hole rubber stoppers, and a section of glass tubing was used as a sleeve to prevent curling of the PTFE tubing and to keep the end of PTFE tubing touching the bottom of the reservoir. Each reservoir was placed on a digital balance (Ohaus SPX2201) with a digital interface (Ohaus Scout RS232 interface) for measurement of the volume (weight) remaining in the reservoir at any given time.
When in turbidostat mode, constant average turbidity was maintained. Specifically, the pump was activated when the measured OD was above the set point and deactivated when the OD was below set point. OD was measured using an 940nm LED (Ledtecch UT188X-81-940, driven with 50ma current) and phototransistor (Ledtech LT959X-91-0125). Each LED-phototransistor pair was tested and selected for consistent OD measurements. The LED and phototransistor were positioned by mounting holes on the aluminum metal frame, on opposite sides of the reactor vessel, 4 cm from the vessel bottom. Each phototransistor was connected to an op-amp (LM324) circuit that acted as a current to voltage converter and buffer (Fig S10B). An isolated DC-DC converter provided a regulated voltage supply for the electronics. The output voltage from the photodetector circuit was digitized using a DAQ (National Instruments USB-6009), and read by the LabView program for OD measurement. The Labview program stored the average light intensity I 0 over the first two minutes after starting a channel as the "blank" value. , The light intensity, I, was measured every ~30s, and the OD = log 10 (I/I 0 ) was calculated.
When in chemostat mode, a constant average flow rate f of medium into the vessel was maintained. Unlike our earlier chemostat setup 47 , here constant flow rate was achieved via a scale (Ohaus SPX2201) which constantly weighed its associated reservoir, and the reading was acquired through an RS232 interface (Ohaus Scout RS232). The initial scale reading, M initial , was recorded when the chemostat channel was activated or reset. This was used with the current scale reading M current to calculate the total volume pumped from the reservoir to the vessel as 1ml/g * (M current -M initial ). The target volume that should have flown from the reservoir to the vessel at the current time was calculated according to the preset flow rate. If the total volume was less than the target, the pump was activated, and otherwise, the pump was deactivated. This provided the correct average flow rate. The flow rate was chosen for the desired doubling time, f = ln(2) * V / T D , where V is the volume of the vessel and T D the doubling time. Vessel volumes were measured by weighing empty vessel, and then weighed again when filled to the spillover point (Fig S10E), giving an average value of 43ml. Individual flow rates were determined using individual vessel volumes. Volume measurements are limited by the minimum waste tube drop size of ~0.5 ml, which is constrained by surface tension. Scale readings were logged, providing a measure of flow rate (Fig S10D).

Bioassays
Most bioassays are yield-based. Various tester strains were used in this study (specified in each experiment; Table S1), but preparation for all yield-based bioassays was the same. Strains were grown for ~16 hours in SD and any required supplements. During this time, growth rate was tracked to ensure cells were doubling as expected (1.6 to 3 hour doubling depending on the strain/condition). After this time, cells were washed 3 times with 3 mL SD and starved for at least 3 hours at 30 o C in 3 mL SD in a never used 13 mm test tube (to prevent inadvertent nutrient contamination). This was done to deplete cells of vacuolar stores of nutrients. Finally, ~1000 cells/well were inoculated into a final volume of 150 µL into a flat-bottomed 96 well plate of either metabolite standards or supernatants, supplemented with SD+1x lysine. For each auxotrophic strain, SD+1x lysine supplemented with various known concentrations glutathione were used to establish a standard curve that related organosulfur concentration (in terms of fmole GSH equivalent) to final turbidity (Fig S4). Turbidity achieved in a supernatant was then used to infer organosulfur concentration in the supernatant. Plates were wrapped with parafilm to prevent evaporation and incubated at 30˚C for 2 days. We re-suspended cells using a Thermo Scientific Teleshake (setting #5 for ~1 min) and read culture turbidity using a BioTek Synergy MX plate reader.
In a rate-based bioassay (Fig S3B), mCherry tagged yeast strain auxotrophic for organosulfur (WY2035) was pre-grown in SD +164 µM Lys +134 µM Met and growth rate was tracked by optical density to ensure the cell was growing as expected. Next, cells were washed 3 times in SD lacking supplements, and starved for at least 3 hours in never-used 13 mm test tubes (to prevent nutrient contamination). Cell density was measured again by OD, and cells were inoculated to roughly 1000 cells /well in a 96 well plate in a total volume of 300 µL. The well was filled with either known quantities of organosulfur (methionine or glutathione) or harvested supernatants, both supplemented into SD + 164 µM lysine. The 96 well plate was measured in the same manner as previously outlined in the "Fluorescence microscopy" section.
Maximal grow rate was calculated by measuring the slope of ln(Normalized Intensity) against time. For each sliding window of 4 time points, slope is calculated and if it exceeds the current max slope for the well, it is chosen as the new maximum. To ensure that no estimation occurs when other metabolites, such as glucose, could be limiting, we restricted analysis to data at 25% maximal intensity to ensure that cells had at least 2 doublings beyond when they are theoretically growing maximally. For this study, maximal growth rates were used to estimate approximate niche size, the logic being that cells should grow faster if there is a larger organosulfur niche.

Flow cytometry
Detailed description can be found elsewhere 48 . Population compositions were measured by flow cytometry using a DxP10 (Cytek). Fluorescent beads of known concentration (as determined by hemocytometer) were added to determine cell densities. A final 1:20,000 dilution of ToPro3 (Molecular Probes T-3605) was used for each sample to determine live and dead cell densities. Analysis using FlowJo software showed obvious clustering of live and dead cells in the ToPro3 RedFL1 channel, with dead cells having a RedFL1 signal of >10 3 . Dead cell densities typically were never higher than 10% in all conditions tested.

Metabolite extraction
Metabolite extraction for intracellular organosulfur quantification was adapted from 49 . Briefly, 20 mL of chemostat populations are harvested in disposable 25 mL pipettes, and rapidly vacuum filtered onto precut 0.45 µm Magna nylon filters (GVS Life Sciences, USA). Filters were then quickly submerged with ethanol cleaned forceps in into 3 mL ice-cold extraction buffer (held in a 5 mL sterile centrifuge tube) mixtures, which consisted of 40% (v/v) acetonitrile, 40% (v/v) methanol, and 20% (v/v) of distilled water. All reagents were HPLC grade and all extraction buffer mixtures were made fresh before each extraction. The centrifuge tube was capped and quickly vortexed to remove all cells. The entire process took less than 25 seconds, with the time between populations being filtered and submerged in extraction buffer being less than 10 seconds. After all populations had been harvested, extracts were frozen at -80 o C until solid, transferred to ice and allowed to thaw. After the samples had thawed, they were incubated on ice for 10 minutes and vortexed once every ~3 minutes and returned to -80 o C for refreezing (a single 'freezethaw' cycle). After 3 freeze-thaw cycles, 1.5 mL of sample was harvested and transferred to a new 1.5 mL micro-centrifuge tube, and centrifuged at 13,0000 rpm for 2 minutes at 4 o C to pellet the cell debris. The extract was removed and the remaining cell pellet was extracted again with 1.5 mL of extraction mixture, and spun down. The final result was 3 mL of extracted metabolites that was stored at -80 o C and analyzed by HPLC less than 48 hours after extraction. To check that a majority of metabolites were extracted, 100 µL of fresh extraction buffer was added to the collected cell debris, vortexed vigorously, and collected by centrifugation. This 100 µL 'second extract' was also analyzed for glutathione by HPLC. On average, the amount of glutathione in the second extract was < 2% of the amount extracted initially in the 3 mL extraction.

HPLC
Reduced glutathione was derivatized using a thiol-specific probe first described by 50 , called Thiol Probe IV (EMD Millipore) to make a fluorescent glutathione conjugate. The compound reacts readily with freethiols, though at different rates. For quantifying glutathione, 270 µL of sample or GSH standard in SD was added to 30 µL of 833 mM HEPES buffer, pH 7.8. This was done to raise the pH of the sample to a basic level, which facilitates the reaction. Next, the probe (dissolved in DMSO and stored in 50 µL aliquots at -20 C), was added to a final concentration of 100 µM, which is in excess of glutathione by at least 10-fold. The reaction was performed at room temperature in the dark (the probe is light-sensitive) in . The SecurityGuard (pre-column) was periodically replaced whenever pressure reading exceeded the manufacturer's specifications. Glutathione was eluted from the column with a mobile phase gradient of filtered Millipore water (Solution A) and acetonitrile (Solution B, HPLC grade). The Millipore water was filtered through a 0.22 µM filter prior to use. Additionally, before each run the column was equilibrated for 30 minutes with 1% Solution B. The % solution B followed the following program for each injection: 0 min 1%, 10 min 14%, 10.01 min 1%, and 15 min 1%, corresponding to a gradual increase to 14% Solution B over 10 minutes, followed by a re-equilibration with 1% Solution B. The column was maintained at a running temperature of 25 o C in a Nexera X2 CTO-20A oven. Flow rate was 1 mL /min. Under these conditions, glutathione eluted at ~ 7 minutes, with slight run-to-run variation. Fluorescent glutathione was detected by excitation at 400 nm and emission at 465 nm. After each run, the column was washed and stored per manufacturer's instructions.
Analysis of HPLC data was done using the R Statistical Language with custom written software for peakpicking, baseline correction, plotting, and area estimation, which is freely available at https://bitbucket.org/robingreen525/hplc_rscripts/src/master/ . Raw data for each sample run was exported to a text file and parsed in the RStudio environment. Emission data (at 465 nm) was culled to restrict analysis from 6.5 to 8 minutes. Next, the script identified a local maximal peak, which for concentrations above 0.01 µM glutathione always corresponding to glutathione. Anything lower was indistinguishable from the background. Next, the script identified local minima on both sides of the glutathione peak and drew a baseline that connected the two. The formula (y=mx+b) for this line was calculated and the baseline was 'corrected' by subtracting the emission spectrum value for each point against the y value of the calculated formula for the same point. To quantify the concentration of glutathione in a sample, known concentrations of GSH in SD were subjected to the above procedure. A standard curve of 0.03 to 1 µM was typically used and showed little variability between experiments. An example of these data are shown in Figure 2A. A linear regression model of peak area against concentration of reduced glutathione was built using the lm function of the stats package. Samples with areas within the dynamic range (0.03-1 µM GSH) were back-calculated using the linear regression model. Comparing HPLC traces of the same derivatized sample over 24 hours shows that glutathione peak area is within 10% of all replicates.

Distinguishing live versus dead release and calculating release rate
To calculate the amount of glutathione each dead cell would need to release for the total pool of glutathione to be explained by cell lysis alone (the 'dead release' model), concentration of supernatant glutathione (in µM) was divided by dead cell density (cell/ mL) as determined by TO-PRO straining (See Flow Cytometry section). This number was multiplied by 1x10 6 to get fmole/dead cell, the amount each dead cell would need to have inside it to explain the total pool of glutathione by cell lysis alone. This method has previously been described by 52 .
To calculate the release rate of glutathione per live cell in chemostats, we used a previously described method 48 . Briefly, steady state concentration of glutathione in chemostat was divided by live cell density as determined by TO-PRO straining, and multiplied by dilution rate (/hr),

Competition
To quantify multiple competition replicates at multiple initial strain ratios, we used the coculture system to mimic the lysine-limited environment, especially since similar mutations in coculture and monoculture lysine-limited chemostats meant that the environments were similar. To do so, WY1340 (the purine requiring/lysine releasing strain in the RM11 background) was grown to exponential phase overnight in SD+134 µM adenine, washed 3 times with SD to remove adenine, and starved for 24 hours to deplete vacuolar storage. During this starvation, WY2702/2073 (BFP met10 evolved clones) and WY2429 (mCherry MET10 evolved clone) was grown overnight in SD + 134 µM methionine to exponential phase, washed 3 times, washed 3 times with SD to remove excess methionine and lysine. Next, WY2072/3 and WY2429 were mixed in ratios of 1:100, 1:10, 1:1, and 10:1 to a final OD 600 of 0.1. This mixture of populations was then added 1:1 with WY1340 to a final OD 600 of 0.03. This was considered generation 0. Populations were monitored for growth by measuring optical density over time and periodically diluted back to OD600 0.03 (OD 600 was never greater than 0.45 to ensure no additional metabolites from SD was limiting). The OD 600 data was used to back calculate total generations in the experiment. Periodically, 100 µL of the culture was sampled for flow cytometry to track strain ratios. Experiments were performed until the strain ratio stabilized.
The steady state species ratio may be estimated in the following fashion. At steady state species ratio, organosulfur releasers and consumers grow at an identical rate, and organosulfur concentration is in steady state. From 8-hr doubling time chemostat experiment (Fig 3), the steady state concentration of organosulfurs was 1.4~1.5 µM GSH equivalent, and the steady state live population size was 4.6~4.8x10 6 /ml. Thus, the release rate was ~0.026 fmole GSH equivalent/cell/hr. From Fig S4 and since in our setup 1 OD~ 3x10 7 cells/ml, organosulfur consumption per birth during starvation was 2.1 fmole GSH/cell. The organosulfur concentration O at steady state can be described as: where R and C are the concentrations of releaser and consumer cells, respectively, and g, r, and c represent growth rate (equal dilution rate in chemostats), release rate of organosulfurs, and organosulfur consumption amount per birth, respectively. Setting this to zero, and we obtain: . This is consistent with Fig 4 where 1. Evolution of organosulfur auxotrophy in cells suffering nutrient-growth imbalance. Fig 1 (A) lys-cells suffer nutrient-growth imbalance when limited for lysine but not when limited for glucose. Exponentially-growing mCherry-expressing lys-(WY2490) cells were washed and starved for lysine and glucose to deplete cellular storage, and cultured and imaged in indicated environments. Total fluorescence serves as an indicator of total live biomass, increasing if cells grow in size or give birth to new cells, and decreasing if cells die 48 . In the absence of lysine, lys-cells died rapidly when glucose was abundant (2%, "Excess C", dark blue circles), and rapid death could be rescued by the TORC1 inhibitor rapamycin (1 µM, blue crosses). lys-cells survived well when the medium lacked glucose in addition to lacking lysine ("No C", green circles). These observations are consistent with the following model: In abundant natural nutrients without lysine, cells attempt to grow, leading to cell death (iii). When carbon is missing (ii) or when rapamycin is applied to inhibit TORC1 (i), cells stop growth, engage in starvationresponsive program, and survive lysine limitation. Total fluorescence intensity is normalized to the initial value, and error bars correspond to 2 standard deviations for 5 replicate wells. (B) The emergence of lys -orgS auxotrophs. lys-cells (WY950 or WY1335) were grown for tens of generations in minimal medium SD containing limiting lysine and no other amino acids, either in lysine-limited chemostats 53 or via coculturing with a lysine releaser in a cross-feeding yeast community 48,54 . Out of 20 independent lines, we randomly isolated ~70 clones for whole-genome sequencing. Chemostat evolution and coculture evolution both yielded met-mutants (1 out of 9 clones in chemostat evolution; 3 out of ~60 clones in coculture evolution). These mutants required an externally-supplied organosulfur such as methionine to grow. In the experiment shown here, clones were grown to exponential phase in SD supplemented with amino acids, washed with SD, starved for 3 hours to deplete cellular storage, and spotted on indicated agar plates for 2 days at 30 o C.  Organosulfur concentration was measured in terms of GSH equivalents using a met10-turbidity-based bioassay (Fig S4C), with brown dotted line marking the lower limit of the linear detection range. we have identified (orange) fail to synthesize homocysteine, and thus can be supported by any of the organosulfurs depicted here as well as potentially other organosulfurs.

Fig S2. lys -orgS repeatedly rose to detectable frequency when lys-cells were limited for lysine
Fig S2 lys-cells (WY1335) were either cocultured (crosses) with a lysine-releasing strain (WY1340) or cultured alone in lysine-limited chemostats (circles). Cultures were frozen periodically in 50% trehalose. We revived frozen samples by directly plating samples on YPD or, in case of cocultures, YPD + hygromycin to select against the partner strain. We observed a variety of colony sizes, and since large and small colonies had different percentages of lys -orgS -, we quantified both types and calculated the overall orgS-in the population (Methods). Colonies that grew in YPD and SD+Lys+Met but failed to grow in SD+Lys were scored as lys -orgS -. Different colors indicate independent evolution lines. The fraction of lys -orgS is much lower in monocultures compared to cocultures. One possibility is that in coculture experiments, the lysine-releasing strain also released organosulfurs. Organosulfurs in supernatants were measured as "methionine equivalents" by comparing growth rates of a met17-tester strain (WY2035) in supernatants fortified by SD versus in SD supplemented with known concentrations of methionine (standard curve in inset). Since the growth rate of tester cells can be affected by factors other than organosulfurs (e.g. pH), the organosulfur niche estimated by a rate-based bioassay can differ from that estimated from a turbidity-based bioassay (e.g. Fig  3B). Nevertheless, this assay motivated the LC-MS experiments that identified GSH as one of the released organosulfurs (Fig 2A). Specifically, the steady state concentration of a released chemical in a chemostat can be expressed as [release rate]*[live cell density]/[dilution rate] 48 . Thus, released chemicals should be twice as concentrated in 8-hr chemostats as in 4-hr chemostats due to the different dilution rates if all else are constant. In this experiment, although the steady state had not been reached by 72 hrs, organosulfurs in 8-hr chemostats were generally ~4-fold those in 4-hr chemostats, consistent with a faster organosulfur release rate in 8-hr chemostats than in 4-hr chemostats (C). Regardless, using liquid chromatography-mass spectrometry and the criterion that steady-state supernatants from 8-hr doubling time chemostats contained more organosulfurs than those from 4-hr doubling time chemostats, we identified glutathione as a released organosulfur (Fig 2A). (C) The release rate of organosulfurs is higher in 8-hr doubling chemostats compared to 4-hr doubling chemostats. Release rates were calculated from the first 24 hrs. A total of five replicates from two experiments are plotted. If organosulfurs are the major released compounds that affect met17 growth rate, then release rates in 8-hr chemostats are significantly higher than in 4-hr chemostats by t-test. . For both gsh1-and met10-strains, since reduction of one GSSG molecule generates two GSH molecules, culture maximal turbidity in GSSG should be twice as much as in equal-molar GSH. This is indeed observed. (E) gsh1-and met10-grew to similar levels in supernatants of lys-cells grown in lysine-limited chemostats. Fig S5 (A, B) Ancestral lys2-cells (WY1335) were cultured in excess lysine (turbidostats with doubling time of 1.5 hrs; magenta) or limited lysine (lysine-limited chemostats at doubling time of 8 hrs; green). Live and dead population densities were quantified using flow cytometry (Methods), and supernatant GSH concentrations were quantified using a fluorescence-based HPLC assay 50 (Methods). Different symbols represent independent experiments. The dotted line in B marks the sensitivity of the HPLC assay.

Fig S5. Lysine limitation increases GSH release rate
(C) Higher GSH release rate during lysine limitation. GSH release rate by lys2-was calculated from the average steady-state GSH concentration, steady-state live cell density, and dilution rate (Methods). Pvalue was derived from a one-tailed t-test assuming equal variance. (D) GSH release rates were comparable between the ancestral lys2-(circles) and an evolved lys2-clone (crosses; WY2429 which contains an ecm21 mutation and Chromosome 14 duplication and thus exhibit improved affinity for lysine). Here, cells were grown in lysine-limited chemostats (8-hr doubling). In (C) and (D), different colors correspond to experiments done on different days. Fig S6 Ancestral (WY1335, circles) and evolved (WY2429, crosses) lys-cells were cultured in lysinelimited chemostats (doubling time 8 hrs). Intracellular metabolites were extracted from cells to quantify fmole GSH/cell (brown). We quantified dead cell density and the concentrations of GSH in culture supernatants. We then calculated the theoretical amount that would need to be inside an average cell in order for cell lysis alone to explain the supernatant concentrations (purple). Since the theoretical amount was higher than the actual amount in all experiments, GSH is likely released by live cells. GSH was quantified using HPLC, and dead cell density was quantified using flow cytometry (Methods). Here, each column corresponds to an experiment. Day-to-day variations exist, but the trend is clear across days. were competed in a lysine-limited environment by coculturing with a lysine-releasing strain (WY1340). Strain ratios over time were measured by flow cytometry (Figure 3). For each trajectory, we computed the slope of ln(lys -orgS -: lys -) over three consecutive time points, and chose the steepest slope. Since our time unit was generation, we divided this slope (/generation) by ln2/generation and obtain a dimensionless number representing the relative fitness difference between lys -orgS and lys -. We then plotted the relative fitness difference against the fraction of lys -orgS at the beginning of the time window used to calculate the steepest slope. Dotted line marks equal fitness between the two strains.

Fig S8. lys -orgS survives lysine limitation better than lys during sulfur limitation
Fig S8 lys -(WY2429, blue), lys -orgS -(WY1604, orange), and lys -orgS -atg5 -(WY2370, green) lysine auxotrophs were grown in SD+ excess lysine (164 µM) and excess GSH (134 µM) to exponential phase. These cells were washed and starved in SD for 5 hours prior to imaging in indicated conditions with (crosses) or without (circles) 1 µM rapamycin in the absence of lysine. Total fluorescence normalized against time zero are plotted. (A) When GSH was limited, lys -orgS survived better than lys -(orange circles above blue circles). TORC1 inhibition by rapamycin improved the survival of both lys -orgS and lys to comparable levels (orange and blue crosses). lys -orgS -atg5 survived poorly even when TORC1 was shut down (green crosses). (B) At high GSH, both lys -orgS and lys survived poorly (orange circles trending down at a similar slope as blue circles), and this poor viability was rescued by rapamycin (blue and orange crosses). lys -orgS -atg5 survived poorly in the presence or absence of rapamycin (green).
Error bars represent two SEM (standard error of mean) from six wells.

Fig S9. Comparing the growth and death profiles of lys orgs and lys cells
Fig S9 lys -(WY2429, blue) and lys orgs -(WY1604, orange) cells were grown in SD+ excess lysine (164 µM) and excess GSH (134 µM) to exponential phase. These cells were washed and starved in YNB+2% (w/v) glucose for 24 hours and imaged in various concentrations of GSH and lysine. During the growth phase, lys grew faster than lys orgs -. After lysine was exhausted, lys orgs survived better than lys -. Note that in the microscopy assay, the minimal medium did not contain GS-X or other excreted compounds found in culture supernatants. Regardless, and consistent with the competition experiment, lys orgs cells grew faster than lys cells under certain conditions. In this experiment, at 2 µM GSH and 0.1 µM lysine, the maximal growth rate achieved by lys orgs was 0.153±0.009/hr (mean ± 2 standard error of mean), greater than the 0.133±0.004/hr achieved by lys -. Fluorescence intensities of various time points are normalized against that at time zero.

Fig S10. Continuous culturing device
The continuous culturing device (A) consists of six channels which can independently operate as a chemostat or turbidostat. Each channel (B) consists of a vessel containing the culture, a magnetic stirrer, an LED-phototransistor optical detector for OD measurement, a computer activated pump, a media reservoir, and a scale for measuring media reservoir (and flow rate). A LabView program running on the CPU uses data from the scale or optical detector to control the pump, and maintain a constant average OD in turbidostat mode, or a constant average flow rate in chemostat mode. Each vessel (C) consists of a Pyrex test tubes modified by adding a waste outlet of adequate diameter and slope to ensure a reliable flow of waste driven by gravity. The vessel's rubber stopper was outfitted with a sampling port consisting of a needle that can be raised and lowered through a seal provided by a segment of PharMed tubing. The tightness of the seal can be adjusted using a Zip-tie, allowing easy motion of the needle, while maintaining its position when stationary. Glass tubing? The six vessels, stirrers, and photodetectors are held in position in a frame cut from an aluminum bar. The signal from each phototransistor was converted to a voltage using an op-amp current to voltage converter (B). (E) Reactor volume averages 43 ml. Individual reactor volume was used for converting doubling time to flow rate, and was measured to ~0.5 ml resolution (limited by minimum outflow drop size; error bar). Table S1. Strains used in this study. Table S2. Mutations in evolved clones Table S2 All nine clones with the alias "ACI" are from chemostat evolution experiment. All other clones are from coculture evolution experiments, a fraction of which (i.e. "CT" strains) are from an earlier study 13 . For WY2467, earlier experiments noted that the strain failed to grow in SD + lysine. We identified it as met-, but its genetic basis is unclear since among identified mutations, none is known to affect methionine biosynthesis. Alternating blue and white shading is to facilitate visualization. Green shading: auxotroph. Orange shading: a prototrophic clone isolated from the same sample as the neighboring auxotrophic clone.