Experimental Evolution

To increase genetic diversity at the onset of the experiment, we founded five experimental (B1–B5) and three control (K1–K3) populations from an F1 cross. We obtained plus (CC-1690) and minus (CC-2290) mating type strains of C. reinhardtii from the Chlamydomonas Resource Center (www.chlamycollection.org) and crossed them using a protocol adapted from Harris20. Vegetative cells were grown to high density in 10 mL TAP medium33 at 22.5 °C and pelleted in 15 mL conical tubes by centrifugation for 5 min. at 2000 × g. Pelleted cells were resuspended in 10 mL mating medium (TAP minus NH 4 Cl) and grown in light for 6 h to induce gametogenesis. Gametes were mixed in a 24-well tissue culture plate with 0.5 mL of each mating type, incubated in light for 4 h, and then dried in the dark. Single mating-type controls of 1 mL were treated identically to ensure that no vegetative material survived desiccation. After 20 days in the dark, crosses and single mating-type controls were each flooded with 1 mL TAP medium and placed in light at 22.5 °C to germinate. Single mating-type controls showed no evidence of vegetative growth after two weeks; thus, desiccation was effective in killing vegetative material and surviving algae from the crosses must be outcrossed.

Experimental populations were started by mixing 0.1 mL of C. reinhardtii F1 cultures, grown to ~2 × 106 cells/mL, with 0.1 mL of a culture of P. tetraurelia, grown to ~2 × 105 cells/mL, in 1.3 mL of COMBO medium34, making a final volume of 1.5 mL. Control populations were founded by inoculating 0.1 mL of C. reinhardtii F1 cultures in 1.4 mL of COMBO medium. All populations were propagated in 24-well plates and transferred weekly.

During transfers, populations were homogenized with a multichannel pipette by drawing and dispensing 1 mL three times. After mixing, 0.1 mL of culture was transferred to 1.4 mL fresh COMBO in a new 24-well plate. After 350 days, populations were plated on TAP medium +1.5% agar. For each population plate, eight individual colonies were randomly picked and re-plated on agar. Individual colonies were picked and re-plated at least three times each to ensure that each was monoclonal and free of contaminants.

We initially characterized evolved phenotypes using light microscopy. A homogenized dilution of each isolate grown in TAP medium was fixed in iodine and stored at 4 °C. For each isolate, 48 µL samples were photographed in replicate on a hemocytometer. Images for each of the clones were processed in ImageJ to determine particle counts and relative particle sizes, and these data were later processed in Python to determine the relative proportions of different sized particles. The size threshold used to distinguish between unicellular and multicellular particles was user-determined and was rarely violated by any samples. In all instances where the size threshold was violated, small multicellular particles were categorized as unicellular; thus, we remain conservative in our characterization of particle sizes, under- rather than overestimating the number of cells in particles.

Maximum Cells Per Cluster

In order to determine the size attained by different evolved strains, measured as the maximum number of cells per cluster, we stained and imaged samples via fluorescent microscopy over a 6-day examination period. Strains were cultured in 10 mL of TAP medium in a climate-controlled chamber on a 14:10-hour light:dark cycle (23.5 °C, 41% humidity, 326 lx). In order to standardize starting cell concentrations, preliminary cultures were first inoculated from scrapes of colonies on TAP plates (2% agar) and allowed to grow, without disruption, for 6 days until reaching stationary phase. Cultures were then transferred 1:100 into fresh TAP medium and examined over the following 6-day examination period.

Throughout the examination period, cultures were sampled, stained, and imaged every 12 hours in order to capture the maximum size attained by various strains before reaching stationary phase. Strains B5-05 and B5-06 were not sampled at the final time-point. Samples were extracted from well-mixed culture tubes and transferred into 1.5 mL microcentrifuge tubes (200 μL from the first 6 time-points to compensate for low cell density, 100 μL from all time-points thereafter). The samples were centrifuged at 250 × g for 5 minutes and the supernatant was replaced by a 50% v/v ethanol-water solution to fix cells and make cell membranes more permeable to the DAPI (4′,6-diamidino-2-phenylindole) fluorescent nucleotide stain. After a 10-minute fixation period, samples were centrifuged again (250 × g for 5 minutes) and the supernatant was replaced with distilled water containing 1 μg/mL DAPI. Cells with DAPI were kept in darkness for one hour and then moved to a refrigerator without light to prevent photo bleaching until imaging on the microscope.

Prior to imaging, samples were removed from the refrigerator and inverted to mix cultures. Seven μL of each sample were mounted on glass microscope slides and large 6 × 6 images were stitched from individual images captured at 40x magnification, creating high-resolution (441 megapixel) wide-field images, providing a robust measurement of the distribution of clusters sizes within the population.

Image and Data Analysis

Large stitched images of individual samples were imported and analyzed in Fiji35. We wrote and implemented a script in ImageJ macro (Supplemental Material) to automatically calculate the number of cells in each cluster. The script accomplished this by demarcating cluster boundaries and recording the number of pixel maxima within each boundary, corresponding to the number of DAPI-stained nuclei. Following this procedure, all images used in our analyses were manually screened and any regions of interest resulting from artifacts (e.g., dust particles, autofluorescence) were removed from the dataset. Data for five technical replicates of all tested strains, each represented by 12 time-points sampled over six days, were concatenated into one large dataset.

In order to determine the maximum number of cells per cluster achieved by each strain, hereafter referred to as “cluster size”, we first determined the time-point at which clusters were largest. This was calculated separately for each strain, as different strains reached their maximum size at different time-points during the six-day culture cycle. First, we calculated the median cluster size of each replicate at each time-point throughout the culture cycle. Medians were chosen to represent the central tendency instead of the mean because the distributions of cluster sizes were right-skewed and occasionally contained large outlier clusters. Then, to determine the time-point at which cluster size was largest for each strain, we calculated the average (mean) of the median cluster sizes from replicate populations within each time-point. The first time-point, representing cluster sizes at time of inoculation, was removed from analysis.

Number of Cells Per Propagule

In order to determine the number of cells per propagule released by clusters of each strain, hereafter referred to as “propagule size”, we time-lapse imaged all strains over 72 hours of growth. First, cells were inoculated from colonies on TAP agar plates into liquid TAP media and incubated for five days to grow to high density. Then, each culture was homogenized by vortexing, transferred 1:100 into five replicate tubes of fresh TAP media, and allowed to grow for two days. These exponentially growing cultures were mixed by vortexing, diluted 1:10 in TAP medium, and then 100 μL from each culture was randomly inoculated into the central 60 wells of a 96-well tissue culture plate that already contained 100 μL of TAP medium in each well.

For time-lapse microscopy, the 96-well plate (containing five technical replicates of each strain) was imaged at 200x magnification, where the field of view was positioned on cells near the center and at the bottom of wells. The time-lapse was run for 72 hours, capturing images of each well every 30 minutes.

Time-lapse images were manually analyzed using ImageJ. For each well, all images from the time-lapse were viewed individually and sequentially. All cells or clusters of cells in the frame at the beginning of the time-lapse were labeled with a number indicating their identity. Throughout the time-lapse, when a cell or cluster of cells was seen to separate from the initial parental cluster/group, the number of cells in the propagule was recorded. For wells that had few or no initial cells in the frame (typically occurring for motile, unicellular strains), cells or clusters of cells were labeled as they entered the field of view, until collecting a sufficient sample size (N > 5 parental clusters). Due to the long time required for clusters to fully separate physically—even after they have detached from a parent cluster—propagule sizes were recorded four frames (two hours) after the propagule was observed to initially split from the parental cluster. Data collected from time-lapse images were analyzed in R36.

Defense Against Predators

To test whether multicellularity affords protection from predators, we subjected evolved unicellular and multicellular isolates to predation by the rotifer Brachionus calyciflorus. Rotifers were chosen for this assay instead of Paramecium to reduce the likelihood that some adaptation other than cluster formation protected cluster-forming strains from the predator. Rotifers were obtained as cysts in vials and stored at 4 °C in the dark. To ensure that cultures of B. calyciflorus were free of contaminants, cysts were treated and incubated according to the protocol described by Suga et al.37. Neonate rotifers were grown axenically in 10 mL of WC medium38 with 100 μg/mL ampicillin in culture tubes at the same temperature and light conditions as the C. reinhardtii cultures. In order to sustain the population of B. calyciflorus before the experiment, 1.0 mL of an axenic, unicellular strain of C. reinhardtii, CC-1690, was inoculated into the culture via micropipette.

Evolved isolates of C. reinhardtii were grown to high density, and culture absorbance was measured using a microplate reader (Molecular Devices SpectraMax® M5, 420 nm), whereafter each was diluted to approximately 3.45 × 105 cells/mL. Four replicates of each strain were randomly assigned to 24-well plates and 1.33 mL of each strain pipetted into designated wells. Twelve multicellular strains (B2-03, 04, 06, 07, and 11; B5-01 through 05, 07, and 08) and twelve unicellular strains (B2-01, 10, and 12; B5-06; K1-01, 02, and 04 through 09) were used for this assay. Calibration curves relating absorbance to cell density were established for each strain by comparing absorbance readings to hemocytometer counts. Because our experimental design compared differences of absorbance readings with and without predators for each strain, it is robust to among strain differences in absorbance per cell. That is, it does not require that equal absorbance readings indicate equal cell densities across strains, only that absorbance readings scale linearly with cell density within each strain.

Rotifer cultures were poured through a 25 μm mesh filter to remove the algae they had been feeding on, then pipetted into a sterile test tube awaiting transfer to the experimental wells (C. reinhardtii cells are ~10 μm in diameter and easily pass through the filter). Depending on the treatment condition, 0.67 mL of either WC medium or WC medium with predators was added to each well. Initial absorbance values were recorded and then plates were stored under lights in an incubator on a 14/10-hour light/dark cycle at 22.5 °C for 5 days. Throughout the 14-hour light period, absorbance readings were taken every 3 hours, providing growth rates with high resolution for all strains.