Comparative Phenotyping of Two Commonly Used Chlamydomonas reinhardtii Background Strains: CC-1690 (21gr) and CC-5325 (The CLiP Mutant Library Background)

The unicellular green alga Chlamydomonas reinhardtii is an excellent model organism to investigate many essential cellular processes in photosynthetic eukaryotes. Two commonly used background strains of Chlamydomonas are CC-1690 and CC-5325. CC-1690, also called 21gr, has been used for the Chlamydomonas genome project and several transcriptome analyses. CC-5325 is the background strain for the Chlamydomonas Library Project (CLiP). Photosynthetic performance in CC-5325 has not been evaluated in comparison with CC-1690. Additionally, CC-5325 is often considered to be cell-wall deficient, although detailed analysis is missing. The circadian rhythms in CC-5325 are also unclear. To fill these knowledge gaps and facilitate the use of the CLiP mutant library for various screens, we performed phenotypic comparisons between CC-1690 and CC-5325. Our results showed that CC-5325 grew faster heterotrophically in dark and equally well in mixotrophic liquid medium as compared to CC-1690. CC-5325 had lower photosynthetic efficiency and was more heat-sensitive than CC-1690. Furthermore, CC-5325 had an intact cell wall which had comparable integrity to that in CC-1690 but appeared to have reduced thickness. Additionally, CC-5325 could perform phototaxis, but could not maintain a sustained circadian rhythm of phototaxis as CC1690 did. Finally, in comparison to CC-1690, CC-5325 had longer cilia in the medium with acetate but slower swimming speed in the medium without nitrogen and acetate. Our results will be useful for researchers in the Chlamydomonas community to choose suitable background strains for mutant analysis and employ the CLiP mutant library for genome-wide mutant screens under appropriate conditions, especially in the areas of photosynthesis, thermotolerance, cell wall, and circadian rhythms.


Introduction
The unicellular green alga Chlamydomonas reinhardtii (Chlamydomonas throughout) is a superior model organism to study important cellular processes in photosynthetic eukaryotes, including but not limited to: photosynthesis [1], cell cycle [2], lipid accumulation [3], To evaluate photosynthetic performance, we imaged room temperature chlorophyll fluorescence in algal spots grown on TAP plates using FluorCam, a chlorophyll fluorescence imager (Photon System Instruments). Both strains had indistinguishable PSII efficiency in dark and light on plates. However, CC-5325 had higher non-photochemical quenching (NPQ) than CC-1690 ( Figure 4a). NPQ is one of the most important photoprotective pathways in photosynthetic organisms, helping dissipate excess light energy and protecting photosynthesis [57,58]. To investigate photosynthetic parameters further, we used liquid algal cultures grown in PBRs for detailed photosynthetic measurements. We performed 77 K chlorophyll fluorescence to check the relative antenna size of PSII and PSI [59]. CC-5325 had smaller PSII antenna size than CC-1690 (Figure 4b,c). By using a customdesigned kinetic spectrophotometer/fluorometer, we showed that CC-5325 had higher PSII maximum efficiency in dark-adapted cells but lower PSII operating efficiency under 400 µmol photons m −2 s −1 light than CC-1690 ( Figure 4d). In response to the same light intensities, both strains had little difference in Q A redox status, except at 200 µmol photons m −2 s −1 light (Figure 4e). By using the PSII% (relative PSII antenna size or percentage of light distributed to PSII) estimated from our 77 K chlorophyll fluorescence data and PSII efficiency measured by room temperature chlorophyll fluorescence, we quantified rates of linear electron flow (LEF) in response to light in these two strains (Figure 4f). CC-1690 had higher LEF than CC-5325 only at 400 µmol photons m −2 s −1 light. However, CC-5325 had much higher NPQ than CC-1690 under all three light intensities tested (Figure 4g). Additionally, by using a Hansatech Chlorolab 2 Clark-type oxygen electrode, we found CC-1690 had higher gross O 2 evolution rates than CC-5325 in response to the same light intensities while both had no differences in respiration rates (Figure 4h,i). Our results suggested that CC-5325 had lower photosynthetic efficiency than CC-1690.

. CC-1690 and CC-5325 had comparable growth rates under mixotrophic conditions in photobioreactors (PBRs). Chlamydomonas cells were grown in turbidostatically controlled PBRs in
Tris-acetate phosphate (TAP) medium at 25 • C with a light intensity of 100 µmol photons m −2 s −1 and constantly bubbling of air. Doubling time (a) and relative growth rates ((b), inverse of doubling time) were calculated based on the exponential increase of OD 680 , which is proportional to total chlorophyll content in unit of µg chlorophyll mL −1 and cell density at 25 • C. Mean ± SE, N = 3 biological replicates. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing the two strains. Not significant, ns.
We harvested algal cultures of CC-1690 and CC-5325 from PBRs, spotted them on agar plates with either TAP (with acetate, the supplied carbon source) or TP (Tris-phosphate medium, without acetate) medium, and grew them under different light and temperature conditions in incubators ( Figure 5). CC-5325 grew better than CC-1690 in the dark with acetate ( Figure 5a). CC-1690 could grow in the dark but appeared to have slowed growth as compared to CC-5325. At 25 • C and 35 • C under 150 µmol photons m −2 s −1 light in incubators, the growth of CC-1690 and CC-5325 was indistinguishable (Figure 5b-e,g). To evaluate the thermotolerance of these two strains under well-controlled temperature conditions in liquid cultures, we exposed the algal cultures to 43 • C for 2 h in PBRs, then spotted the cells on agar plates to measure viability. CC-5325 barely survived the heat of 43 • C, while CC-1690 could survive 43 • C heat better than CC-5325 ( Figure 5f). Our results showed that both strains grew well photoautotropically and mixotrophically, but CC-5325 grew better under heterotrophic conditions while CC-1690 had higher thermotolerance than CC-5325.
Thermotolerance is linked to cell wall properties [60,61], although how cell walls affect thermotolerance is unclear. Because CC-5325 is a progeny from a cross between 4A − (cell wall intact) and D66 + (cell wall deficient), CC-5325 is often thought to be cell wall deficient. We evaluated the cell walls of CC-1690 and CC-5325 based on freezing tolerance, a detergent assay, and cell wall imaging (Figures 6-8). Cell-wall deficient strains are thought to recover better from liquid nitrogen freezing than cell-wall intact strains [62]. Both CC-1690 and CC-5325 recovered equally well from liquid nitrogen freezing ( Figure 6). Next, we used a detergent assay to evaluate the integrity of the cell wall. Triton X-100 is a detergent that disrupts lipid-based membranes and causes cells without an intact cell wall to lyse. Strains without an intact cell wall or reduced cell wall integrity will have more damaged cells in the presence of Triton X-100, thus releasing more chlorophyll into the supernatant, which could be quantified using absorbance at 435 nm. We included the well-studied cell-wall-deficient line cw15 as a control for this assay [55,56]. Without Triton X-100 treatment, there was little chlorophyll released in all three lines (Figure 7a). With Triton X-100 treatment, all three lines had increased chlorophyll release, but CC-5325 had the least chlorophyll release while cw15 had the most, demonstrating the effectiveness of our assay. By using a Coulter Counter, we quantified the number of intact cells with and without Triton X-100 treatment (Figure 7b). While Triton X-100 did not affect CC-1690 or CC-5325, it damaged nearly all cells of cw15. Our freezing and detergent assays showed that CC-1690 and CC-5325 had comparable cell wall integrity.

Figure 2. CC-1690 and CC-5325 had comparable growth rates under mixotrophic conditions in photobioreactors (PBRs). Chlamydomonas cells were grown in turbidostatically controlled PBRs in
Tris-acetate phosphate (TAP) medium at 25 °C with a light intensity of 100 μmol photons m −2 s −1 and constantly bubbling of air. Doubling time (a) and relative growth rates ((b), inverse of doubling time) were calculated based on the exponential increase of OD680, which is proportional to total chlorophyll content in unit of μg chlorophyll mL −1 and cell density at 25 °C. Mean ± SE, N = 3 biological replicates. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing the two strains. Not significant, ns.  To evaluate photosynthetic performance, we imaged room temperature chlorophyll fluorescence in algal spots grown on TAP plates using FluorCam, a chlorophyll fluorescence imager (Photon System Instruments). Both strains had indistinguishable PSII efficiency in dark and light on plates. However, CC-5325 had higher non-photochemical quenching (NPQ) than CC-1690 ( Figure 4a). NPQ is one of the most important photoprotective pathways in photosynthetic organisms, helping dissipate excess light energy and protecting photosynthesis [57,58]. To investigate photosynthetic parameters further, we used liquid algal cultures grown in PBRs for detailed photosynthetic measurements. We (f) Chlorophyll/carotenoid ratio. Mean ± SE, N = 3 biological replicates. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing with CC-1690: *, p < 0.05; ns, not significant. To directly visualize and quantify the cell wall, we used the lectin Concanavalin A as a cell wall stain and Syto 13 TM Green Fluorescent Nucleic Acid to label the nucleus, then imaged the cells using a ZEISS Elyra 7 super-resolution microscope ( Figure 8). Our results showed that CC-1690 and CC-5325 had continuous and intact cell walls, while cw15 has discontinuous and broken cell walls, as expected for a cell-wall deficient mutant (Figure 8a-f). We calculated the total cell wall fluorescence intensity along each cell and normalized it to the cell perimeter length. The mean cell wall fluorescence intensity of CC-5325 was 84% of that in CC-1690, while that in cw15 was 17% relative to CC-1690 ( Figure 8g). Based on our results of freezing recovery, detergent assays, and cell wall imaging, we conclude that CC-5325 is not as cell-wall deficient as previously described. Instead, CC-5325 has an intact cell wall with comparable integrity to CC-1690, but the cell wall in CC-5325 is possibly thinner than that of CC-1690.
Next, we investigated the circadian rhythm of phototaxis in CC-1690 and CC-5325 because both phototaxis and circadian rhythms are important for photosynthesis, thermotolerance, and cell growth [63][64][65][66]. Rhythmic phototaxis (swimming towards light) in constant light or darkness is a known circadian-regulated activity in Chlamydomonas and can serve as a read-out of clock function [67,68]. Circadian phototaxis was determined in day/night entrained cultures followed by either "constant darkness" or "constant light" free-running conditions (Figure 9a When monitoring the phototaxis of CC-5325, there seemed a start of a rhythm in nearly all cultures tested in constant light and in most of the cultures tested in constant darkness but only for the first 12 to 36 h of constant conditions. The lack of a sustained circadian rhythm of phototaxis in CC-5325 was independent of cell densities (from 0.5 × 10 6 to 1.6 × 10 6 cells mL −1 ). Our results clearly demonstrated that CC-1690 could maintain a sustained circadian rhythm of phototaxis but CC-5325 could not. of growth. Plate images (all 1:20 dilution) were taken after 10-day (a) or 3-days (b-f) of growth using a regular camera. Images shown are representative of three biological replicates. (g) Normalized cell area. The 44-h spotting images were analyzed by ImageJ to get cell areas which were then normalized to the number of cells spotted. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing with CC-1690 under the same experimental conditions. Not significant, ns. Such quantification was not performed for (a,f) because the cell areas for one of the strains were too small to quantify.  replicates. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing the two strains under the same experimental conditions. No significant differences between these two strains under the same conditions; ns, not significant, p > 0.05. of growth. Plate images (all 1:20 dilution) were taken after 10-day (a) or 3-days (b-f) of growth using a regular camera. Images shown are representative of three biological replicates. (g) Normalized cell area. The 44-h spotting images were analyzed by ImageJ to get cell areas which were then normalized to the number of cells spotted. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing with CC-1690 under the same experimental conditions. Not significant, ns. Such quantification was not performed for (a,f) because the cell areas for one of the strains were too small to quantify.  . CC-1690 and CC-5325 had comparable resistance to the detergent Triton X-100. Detergent assay was performed to check cell wall integrity. Triton X-100 is a detergent to disrupt lipid-based membranes and cause cells without an intact cell wall to lyse. Algal samples with and without 0.05% Triton X-100 were vortexed vigorously. Strains without intact cell walls will have more broken cells in the presence of Triton X-100 and thus more released chlorophyll than strains with intact cell walls. (a) Released chlorophyll from damaged cells was quantified using absorbance at 435 nm. (b) The percentage of intact cells was quantified using a Coulter Counter. Mean ± SE, N = 3 biological replicates. Statistical analyses were performed using a two-tailed t-test assuming unequal variance by comparing with CC-1690 under the same experimental conditions (*, p < 0.05) or within the same strain in the presence or absence of Triton X-100 (#, p < 0.05; ns, not significant, p > 0.05).
8a-f). We calculated the total cell wall fluorescence intensity along each cell and normalized it to the cell perimeter length. The mean cell wall fluorescence intensity of CC-5325 was 84% of that in CC-1690, while that in cw15 was 17% relative to CC-1690 (Figure 8g). Based on our results of freezing recovery, detergent assays, and cell wall imaging, we conclude that CC-5325 is not as cell-wall deficient as previously described. Instead, CC-5325 has an intact cell wall with comparable integrity to CC-1690, but the cell wall in CC-5325 is possibly thinner than that of CC-1690. Because cilia affect phototaxis, we further investigated cilia morphology and swimming speed. Cells were grown in either TAP (with acetate) or M-N (without acetate and nitrogen) medium with constant agitation and light. The M-N condition induces gamete formation and eliminates the effect of the cell cycle on percent ciliation and ciliary length. The percent of cells with cilia was comparable between CC-1690 and CC-5325 in both TAP and M-N medium (Figure 10a,b). However, CC-5325 had significantly longer cilia than CC-1690 in TAP medium; this trend might be also true for cells in M-N medium, but the differences were not significant (Figure 10c,d). We quantified swimming speed by creating traces of how far the cells moved during a 1 s exposure using a darkfield microscopy (Figure 10e

Discussion
We compared phenotypic differences between CC-1690 and CC-5325 by focusing on characteristics that are important for algal growth, including cell size, growth in liquid and on agar plates, photosynthesis, thermotolerance, cell wall integrity, circadian rhythm of phototaxis, ciliary percentage/length, and swimming speed. These two frequently used background strains displayed notable differences in these characteristics ( Figure 11).
Both strains grew well in liquid medium and on agar plates under normal conditions (Figures 2 and 5). CC-5325 grew more robustly in dark with acetate than CC-1690 (Figure 5a), making it a promising background strain for mutants deficient in photosynthesis, which are often maintained in dark conditions with supplied carbon source. The reduced photosynthetic efficiency in CC-5325 may be related to smaller PSII antenna size and/or increased NPQ as compared to CC-1690 (Figure 4c,g,h). The NPQ differences in these two strains may be relative, since NPQ from different strains may not be compared directly [69,70]. The increased NPQ in CC-5325 may be related to the reduced chlorophyll/carotenoid ratio (more carotenoids) (Figure 3f). Carotenoids have reported roles in photoprotection and one of the carotenoid pigments, zeaxanthin, has major roles in NPQ formation [4,71]. The bigger PSI peak of 77 K chlorophyll fluorescence in CC-5325 (Figure 4b,c) suggested it had larger PSI antenna size but reduced PSII antenna size as compared to CC-1690 [59]. CC-5325 has lower oxygen evolution rates than CC-1690 only under light intensity equal to or greater 200 µmol photons m −2 s −1 light (Figure 4h), which may explain the comparable growth rates of these two strains in PBRs under 100 µmol photons m −2 s −1 light (Figure 2). Although CC-5325 had lower photosynthetic efficiency than CC-1690, it does not compromise its suitability as a background strain to screen for mutants deficient in photosynthesis. Because photosynthetic efficiencies are relative, it is not necessary for a background strain to have the highest photosynthetic efficiency for generating and characterizing photosynthetic mutants; normal photosynthesis would be adequate. Previous research showed that CC-5325 has a functional carbon concentrating mechanism (CCM) and several CLiP mutants deficient in CCM have been identified by using CC-5325 as a background strain [25,49,72]. CLiP mutants deficient in photosynthetic light reactions were also identified and published [48,73]. Additionally, a genome-wide mutant screen has been performed to identify genes required for photosynthesis using the CLiP mutant library [38]. Based on our results and published literature, we believe CC-5325 is a suitable background strain to study photosynthesis in Chlamydomonas. CC-5325 has been frequently reported as a cell-wall deficient line, like cw15 [37,50]. Our results from freezing and detergent assays showed that the cell wall integrity in CC-5325 was comparable to that in CC-1690 (Figures 6 and 7). Cell wall imaging and quantification showed that CC-5325 had an intact cell wall, albeit possibly thinner cell wall than that in CC-1690 (Figure 8). The vegetative cell wall of Chlamydomonas is mainly comprised of proteins [74,75]. It is possible that these two strains have different quantities of cell wall proteins. There are three classifications of cell-wall-deficient mutants: (A) normal

Increased
Gross O 2 evolution rates Smaller at or above 200 μmol photons m -2 s -1 light CC-5325 has been frequently reported as a cell-wall deficient line, like cw15 [37,50]. Our results from freezing and detergent assays showed that the cell wall integrity in CC-5325 was comparable to that in CC-1690 (Figures 6 and 7). Cell wall imaging and quantification showed that CC-5325 had an intact cell wall, albeit possibly thinner cell wall than that in CC-1690 ( Figure 8). The vegetative cell wall of Chlamydomonas is mainly comprised of proteins [74,75]. It is possible that these two strains have different quantities of cell wall proteins. There are three classifications of cell-wall-deficient mutants: (A) normal quality of cell walls that are unattached to the plasma membrane; (B) normal quality of cell walls that are attached to the plasma membrane; (C) minimal amount of cell walls [55]. The well-studied cell-wall-deficient mutant, cw15, belongs to classification C [55,75]. Our results showed that CC-5325 was clearly not a C-type cell-wall mutant. Chlamydomonas cell walls protect cells from environmental stresses [75], but the underlying mechanisms are unclear. Further investigation is needed to investigate if the reduced cell wall in CC-5325 contributes to some of its increased heat-sensitivity in comparison to CC-1690 (Figure 5f). Nevertheless, the cell wall integrity in CC-5325 is clearly much better than previously thought and its increased heat-sensitivity as compared to CC-1690 does not affect its use for studying heat responses. The CLiP mutant library has been used to screen for heat-sensitive mutants [54]. It may not be necessary to cross CLiP mutants on the background of CC-5325 to other cell-wall intact strains. Additionally, for research related to heat stress, it may generate conflicting results if the heat-sensitive CLiP mutants are crossed to other non-isogeneic lines (e.g., CC-1690) because different background strains may have different heat sensitivities, as demonstrated in our research. If crossing is needed, a potentially better option would be to cross CLiP mutants to the isogenic strain of CC-5325, which is CC-5155 (mt + ). CC-5155 was generated by crossing CC-5325 (mt − ) and CC-125 (mt + , 137c) and then backcrossing a mt + progeny to CC-5325 five times [37].

Ciliary length Longer in TAP
The most surprising result may be the lack of a sustained circadian rhythm of phototaxis in CC-5325, although the strain can perform phototaxis (Figure 9). Our results clearly showed a sustained circadian rhythm of phototaxis in CC-1690, but CC-5325 only showed a brief rhythm of phototaxis in constant conditions for the first 12 to 36 h. This lack of a sustained circadian rhythm of phototaxis in CC-5325 was present under both constant light and constant darkness. A number of different circadian rhythms have been studied in Chlamydomonas [63,64], including phototaxis, chemotaxis, cell division, UV sensitivities, and others. How well Chlamydomonas can maintain these rhythms under various environmental conditions is less known. From our data, we cannot distinguish whether the lack of a sustained rhythm in CC-5325 is due to defects in the circadian oscillator mechanism, and thereby also affecting other circadian rhythms, or whether it is specifically due to the ability of the clock to regulate specific outputs such as phototaxis. If the former is true, the reduced photosynthetic efficiency and thermotolerance observed in CC-5325 may be related to its deficiency in maintaining circadian rhythms, as normal circadian rhythms are important for photosynthetic organisms to optimize photosynthesis and acclimate to stressful environments [65,66,76,77]. In CC-1690, constant darkness or light affected the period of the rhythm, with a longer period in constant darkness (27.65 h) than constant light (23.61 h). Differences in period length under different constant conditions have been observed in many organisms. For example, it was recently reported that the duckweed, Lemna gibba, showed an average period under conditions of constant darkness that was about 7 h longer than it was in constant light [78]. Transcriptome analysis in cultures under synchronizing versus free-running conditions may provide some insights about the mechanisms of the maintenance of circadian rhythms and about the basis for the observed differences between these two strains.
CC-1690 and CC-5325 also had some differences in cilia length and swimming speed ( Figure 10). CC-5325 had longer cilia in TAP medium than CC-1690, consistent with the increased cell volume of CC-5325 in TAP as compared to CC-1690 (Figure 1), as ciliary length scales with cell size [79]. CC-5325 had slower swimming speed in M-N medium than CC-1690, which may be related to additional influences by positive gravitaxis (downward swimming in response to gravity), as we observed that CC-5325 gravitaxed and settled faster than CC-1690 in both TAP and M-N medium. This may be due to shape-based mechanical orientation of cells [80] rather than due to the large size of CC-5325 as another large-size strain, CC-125, does not exhibit positive gravitaxis. Additionally, the tendency to swim downward did not lead to increased clumping in CC-5325 in our algal cultivation in flasks or algal photobioreactors.
The underlying genetic causes of these phenotypic differences are of great interest. Comparison between genome sequences of CC-1690 and CC-5325 showed that these two strains had 117,059 small variants (include single nucleotide substitutions and small insertions and deletions) and 355 high impact small variants (that are predicted to cause a frameshift or premature translation stop) [37], providing potential candidates for phenotyping differences. To understand the genetic factors underlying the phenotypic differences observed between CC-1690 and CC-5325, we investigated the functional annotations of the nonsynonymous mutations in coding regions of CC-5325 (Supplementary Table S1). Li et al., 2016 reported the high confidence genomic differences that are unique to CC-5325 as compared to several other commonly used laboratory strains, including CC-1690. Among these, we identified the nonsynonymous genetic difference in coding regions of 40 genes in CC-5325, including codon changes, codon insertions/deletions, and frameshift mutations (Supplementary Table S1). Of the 40 genes containing nonsynonymous mutations, six have putative roles in regulation including three transcription factors, three transcriptional regulators, and a histidine kinase response regulator. About half of these genes have no or little function annotation. Notably, we did not observe nonsynonymous mutations in the coding regions of genes with known roles involved in photosynthesis, heat responses, cell wall biogenesis, circadian regulation, phototaxis, and cilia biology. Because CC-5325 had reduced PSII antenna size as compared to CC-1690 (Figure 4b,c), we also checked genes with known roles in state transition in Chlamydomonas. State transition is a process that reorganizes light-harvesting antenna systems and redistributes the light excitation energy between PSII and PSI to balance the energy input/consumption between the two photosystems [81]. The kinase STATE TRANSITION7 (STT7) in Chlamydomonas phosphorylates components of light-harvesting antenna complex II (LHCII), triggering the state transition [81]. Two phosphatases reverse the LHCII phosphorylation and the state transition: PPH and PBCP (Phosphoprotein Phosphatase 2C-related and PSII Core Phosphatase) [73]. However, none of these genes with known roles in state transition had nonsynonymous mutations in CC-5325. These results suggest that the observed phenotypic differences between the two background strains may be due to changes at the regulatory level rather than codon changes in the proteins with known functions involved in these biological processes. Further detailed analysis, such as quantitative trait locus (QTL) mapping, could be performed in the future to identify the causative genetic loci underpinning the phenotype differences observed between CC-1690 and CC-5325. QTL mapping has been used frequently to identify causative genes related to a phenotype in land plants [82][83][84][85]. Such analysis is much less developed in Chlamydomonas. However, Chlamydomonas has several advantages for QTL mapping, e.g., high mating efficiency and easy mating procedure [86], fast growth on plates or in liquid, low cost in whole-genome sequencing [87,88], and abundant RNA-Seq datasets [11]. One round of mating in Chlamydomonas only takes about 2-3 weeks, from the mating process to identifiable progenies. Due to its unicellular nature, high throughput phenotyping tools can be applied to characterize progenies of Chlamydomonas efficiently, e.g., whole plate chlorophyll fluorescence imaging [89][90][91], growth quantification on plates using ImageJ [23], and circadian rhythm monitoring using an automated phototaxis machine [92]. The CLiP mutant library allows for both forward and reserve genetic screens and candidate genes identified by QTL in Chlamydomonas can be validated using available mutants in the CLiP mutant library [37]. If mutants deficient in some specific genes are unavailable in the CLiP mutant library or other mutant collections at the Chlamydomonas Resource Center, mutants disrupted in genes of interest could be generated by CRISPR-Cas9 approaches with high efficiency [34][35][36]. Recently, research conducted to phenotype and select progenies from a cross between two Chlamydomonas parental strains under different environmental conditions revealed possible selection-enriched genomic loci for improved stress adaptation and photosynthetic efficiency [93], paving the way for QTL mapping in Chlamydomonas.

Conclusions
We presented several distinct phenotypes between two commonly used laboratory background Chlamydomonas strains: CC-1690 (21gr) and CC-5325 (CLiP library background). Although CC-5325 has lower photosynthetic efficiency and thermotolerance than CC-1690 and these two strains have some differences in cilia, it does not compromise the use of CC-5325 and the CLiP mutant library to study and screen for mutants deficient in photosynthesis, heat responses, or cilia biology. Our results showed that CC-5325 had an intact cell wall with comparable integrity as compared to CC-1690, which will help reduce researchers' concerns or confusion about its cell wall deficiency. The lack of a sustained circadian rhythm of phototaxis in CC-5325 does limit the use of the CLiP library to screen for mutants deficient in circadian rhythms. There may be no background strains that are perfect for all biological processes, but it is important to understand the phenotypic behaviors of one background strain when using it. Our research will help fill the knowledge gaps about CC-5325 and facilitate the use of the CLiP mutant library generated in this background. Finally, the rich genomic and genetic resources in Chlamydomonas will enable identification of responsible genes for phenotypes related to cell size, photosynthesis, thermotolerance, circadian rhythms, and cilia biology.

Strains and Culture Conditions
Chlamydomonas wildtype strain CC-1690 (also called 21gr, mt + ) and CC-5325 (also called CMJ030, identical to CC-4533, mt − , CLiP library background) were ordered from the Chlamydomonas Resource Center (University of Minnesota). Algal cultures of these two strains were maintained in photobioreactors (PBRs) as described before [23]. The normal growth condition in PBRs was: 25 • C, Tris-acetate-phosphate (TAP) medium, constant 100 µmol photons m −2 s −1 light (50% red: 50% blue), constant bubbling with filtered air (0.04% CO 2 ), and turbidostatic control by monitoring OD 680 to supply fresh medium frequently and maintain cell density around 2 × 10 6 cells mL −1 . Under the normal growth condition at 25 • C, OD 680 is linearly proportional to both chlorophyll content and cell density. Fresh medium was added to the culture automatically by a peristaltic pump when the OD 680 reached the defined maximum value to dilute the culture; the pump was stopped when the OD 680 dropped to the defined minimum value. Algal cultures then grew up at approximately exponential rate to the defined maximum OD 680 value before the next dilution cycle. Our OD 680 range was sufficiently small that we expected minimal nutrient limitation during our experiment. Thus, the turbidostatic mode precisely controlled the growth condition. We calculated the doubling time or relative growth rates based on the exponential growth phase when the peristaltic pump was off. The doubling time and relative growth rates we referred to were based on the increase of OD 680 or the total chlorophyll per mL culture. The growth rates measured by OD 680 were consistent with medium consumption rates and cell number increase quantified by a cell counter. For the 43 • C heat treatment in PBRs, the PBR temperature was gradually increased from 25 • C to 43 • C within 30 min, and then the PBR temperature was maintained at 43 • C for 2 h. By the end of 2 h heating at 43 • C, algal cultures were harvested for spotting test to evaluate viability.

Algal Spotting Test
Algal spotting test was performed as described before with minor modifications [23]. CC-1690 and CC-5325 cultures harvested from PBRs were diluted to 1 × 10 5 cells mL −1 or 0.2 × 10 5 cells mL −1 (1:20 or 1:100 dilution) and 10 µL aliquots of the diluted cultures (about 1000 or 200 cells) were spotted on 1.5% TAP agar plates and grown in temperature-controlled incubators under the indicated conditions. For conditions with light, 150 µmol photons m −2 s −1 light was provided by white LED lights with indicated intensity. After 44-h growth under light in incubators, algal spots were imaged by a dissecting Leica microscope. The whole plates with algal spots were imaged again using a regular camera after 3-day-growth for visual representations. Due to the slow growth in darkness, the algal spots and the whole plates were images after 5 days and 10 days, respectively.

Photosynthetic Measurements
CC-1690 and CC-5325 cultures grown at 25 • C in PBRs as mentioned above were used for all photosynthetic measurements. Algal cultures were spotted on TAP plates and grown under 150 µmol photons m −2 s −1 light at 25 • C in incubators for 4-day. Algal plates were dark-adapted for 20 min before chlorophyll fluorescence imaging at room temperature using FluorCam (Photo System Instruments). Photosynthetic measurements in liquid algal cultures were performed as described before [23]. 77 K chlorophyll fluorescence measurement were performed using an Ocean Optics spectrometer (Cat. No. OCEAN-HDX-XR, Ocean Insight, Seminole, FL, USA) and 430 nm excitation provided by light emitting diodes (LEDs). Spectral data were normalized to the PSII spectral maximum value at 686 nm and relative percentage of light distributed to PSII (or relative PSII antenna size, PSII%) was calculated using this formula: PSII% = normalized PSII peak/(normalized PSII peak + normalized PSI peak at 714 nm). Normalized PSII peak equals one. Room temperature chlorophyll fluorescence in liquid algal cultures were performed using a multiwavelength kinetic spectrophotometer or fluorometer (measuring beam with 505 nm peak emission, measuring pulses of 100 µs duration) [23]. Aliquots of 2.5 mL algal cultures (around 12 µg chlorophyll) were sampled from PBRs, supplemented with 25 µL of fresh 0.5 M NaHCO 3 in a fluorometry cuvette (C0918, Sigma-Aldrich, St. Louis, MO, USA) with constant stirring, and dark adapted with a 10-min exposure to far-red light (peak emission of 730 nm at~35 µmol photons m −2 s −1 ) before chlorophyll fluorescence measurement in dark, and subsequent actinic light phases of 100, 200, 400 µmol photons m −2 s −1 light. The actinic light was provided by LED lights with maximal emission at 620 nm. Each light phase was about 7.5 min long. A saturating flash was applied at the end of each phase to get maximum chlorophyll fluorescence (F m in dark or F m in light). Photosynthetic parameters were calculated as described [70]. PSII efficiency (ΦPSII) was calculated as 1 − F o /F m and 1 − F /F m for dark-adapted and light-adapted algal cells, respectively. F o and F m are minimal and maximal chlorophyll fluorescence in dark-adapted algal cells. F and F m are steady state and maximal chlorophyll fluorescence in light-adapted algal cells. Q A redox status was calculated as fraction of open PSII centres and F q is the photochemical quenching of fluorescence (Baker et al., 2007). Linear electron flow (LEF) was calculated using the formula: LEF = (actinic light) × (Qabs fs ) × (PSII%) × (ΦPSII). Qabs fs is fraction of absorbed light, assuming 0.8. PSII% was calculated from 77 K chlorophyll fluorescence as mentioned above. Non-photochemical quenching, NPQ, was calculated as (F m − F m )/F m . Oxygen evolution was measured using the Hansatech Chlorolab 2 based on a Clark-type oxygen electrode at room temperature as described [23]. Two-mL of cells (around 10 µg chlorophyll) supplemented with 20 µL of 0.5 M NaHCO 3 were incubated in the dark for 10 min with stirring before the light phases of 100, 200, 400 µmol photons m −2 s −1 light. Each light Plants 2022, 11, 585 20 of 26 lasted 5 min followed by 2 min dark. The rates of oxygen evolution and respiration were measured at the end of each light and dark phase, respectively.

Algal Cryogenic Freezing and Thawing
CC-1690 and CC-5325 cultures around a cell density of 2 × 10 6 cells mL −1 were harvested from PBRs grown at 25 • C and frozen in 10% cryoprotective agent (CPA, 1:10 volume ratio of 100% methanol and fresh TAP medium). Algal cultures of 450 µL was added to 450 µL of 10% CPA (final CPA concentration was 5%) in cryo-tubes, mixed gently, transferred to a CoolCell foam freezer container, incubated in a −80 • C freezer for 4 h, and finally stored in a liquid nitrogen freezer. During the thawing process, cryo-tubes with algal samples were thawed in a 37 • C water bath for 5 min and centrifuged at 1000× g for 5 min at room temperature to remove the supernatant. Fresh TAP medium of 900 µL was added to resuspend cell pellets, followed by dark storage of the cryo-tubes without agitation at room temperature overnight. The next day, the cryo-tubes were centrifuged again at 1000× g for 5 min at room temperature to remove the supernatant. Cell pellets were resuspended with 200 µL of fresh TAP medium and 10 µL of the resuspended algal cultures were used for the spotting test to evaluate viability as mentioned above.

Detergent Assay for Cell Wall Integrity
CC-1690 and CC-5325 cultures around a cell density of 2 × 10 6 cells mL −1 were harvested from PBRs grown at 25 • C as mentioned above. A cell-wall deficient strain, cw15 [55], was received from the Umen laboratory at the Donald Danforth Plant Science Center and used as a control. cw15 could not grow well in PBRs probably because its cell wall deficiency made it sensitive to the constant air bubbling in PBRs. So shaker cultures of cw15 around the similar cell density were used for the detergent assay. Growth condition of cw15 on the shaker was 25 • C, around 100 µmol photons m −2 s −1 light, and in TAP medium. One-mL algal cultures were used for chlorophyll quantification [94] and another 1-mL cultures (with 0.05% Triton X-100 or with TAP medium) were immediately vortexed vigorously for 30 s then incubated in dark for 10 min. Then, 500 µL of each sample was withdrawn and centrifuged at 13,000× g to remove cell debris. The absorbance of the supernatant was measured at 435 nm and normalized to chlorophyll contents. The number of intact cells were measured using a Coulter Counter (Multisizer 3, Beckman Counter, Brea, CA, USA).

Cell Wall Staining and Quantification
Ten mL cultures of CC-1690, CC-5325, and cw15 cells as mentioned above were collected and concentrated to 2 × 10 7 cells mL −1 by centrifugation at 1500× g for 5 min at room temperature. Cells were stained with Concanavalin A (ConA, Alexa Fluor ® 594 conjugate) (100 µg mL −1 , ThermoFisher Scientific) [96] and Syto 13 Green Fluorescent Nucleic Acid Stain (5 µM mL −1 , ThermoFisher Scientific, Waltham, MA, USA) for 30 min at room temperature. Stained cells were washed first and then fixed in 4% paraformaldehyde (EM Science, Hatfield, PA, USA) and followed by 1X rinse in TAP medium. Fluorescence images were acquired using a 40X C-Aprochromat objective lens (numerical aperture 1.2) in apotome mode with a ZEISS Elyra 7 super-resolution microscope (ZEISS, Oberkochen, Germany). The 488 nm and 561 nm laser lines with BP 420-480 + 495-550 and BP 570-620 + LP 655 emission filters were used for Syto 13 and ConA, respectively in fast frame sequential mode at 50 ms exposure using dual pco.edge 4.2 sCMOS detectors (PCO AG, Kelheim, Germany). All two-dimensional (2D) images (with 5 phases) were acquired at 49 nm x-y pixel resolution and reconstructed using the SIM 2 module and 2D+ processing with standard live sharpness in ZEN Black 3.0 SR FP2 software. For each strain, 100 cells were quantified by free hand drawing a line intensity profile along the cell wall. The fluorescence intensity of each cell wall periphery was normalized by its perimeter length.

Circadian Rhythm of Phototaxis
CC-1690 and CC-5325 were grown in 125-mL Erlenmeyer flasks containing 50-mL photoautotrophic Tris-phosphate (TP) minimum medium without acetate. Cultures were grown on a shaker at 17 to 19 • C under synchronizing 12 h light/12 h dark cycles. The light intensity during the light phase was about 20 µmol photons m −2 s −1 . When cultures reached cell concentrations between 3.5 × 10 5 and 2.3 × 10 6 cells mL −1 , culture samples were placed into small Petri dishes and their phototaxis was monitored at 20 • C as described [92] with the following modifications. A box around the phototaxis machine with a heating element and ventilators allowed the external temperature around the machine to be kept at a constant but slightly elevated temperature of 24 • C compared to the machine. Additionally, temperature recorders (Tempo Disc, BlueMaestro, The Woodlands, TX, USA) that fit into the Petri dish slots allowed for a more precise monitoring of the temperature the culture samples were exposed to. An algorithm as described before was used to analyze the phototaxis data for their periods [92]. Three independent experiments were conducted for each of the constant darkness and constant light conditions with 1 or 2 different cultures per strain per experiment and 3 to 4 replicate samples per culture.

Ciliary Measurements and Swimming Speed Assay
CC-5325 and CC-1690 cells were grown overnight with constant agitation and light at 100 µmol photons m −2 s −1 light in either liquid TAP or M-N (minimal medium without acetate and nitrogen) to induce gamete formation and eliminate the effects of the cell cycle on cilia length. A sample of each strain was fixed by mixing 1:1 with 2% glutaraldehyde for a final concentration of 1% glutaraldehyde. After fixation, cells were imaged with a DIC microscope. Images were analyzed in ImageJ/FIJI using the segmented line tool. Each cell has two cilia with the same length. One cilium per cell was traced from base to tip, then fit using the "Fit Spline" function [97]. The length of the line was then measured and converted to micrometers. Cells were only included in the measurement if both cilia were visible. The ciliation percentage was obtained by counting the number of cells with at least 1 visible cilium out of 100 cells.
For swimming speed assay, cells were allowed to settle for 15 min so that swimmers could be selected. The swimmers taken from the top of the cultures were diluted 1:10 in the same media they were grown in (either TAP or M-N). To give the cells appropriate room to swim between the slide and coverslip, a chamber was created on the slides using double stick tape. Cells were placed in the chamber and a coverslip was placed on top. Samples were imaged using a darkfield microscope set to a 1-s exposure. The resulting images show traces representing where and how far the cells swam during the 1-s exposure [98]. To estimate the swimming speed, the traces were measured in ImageJ/FIJI using the segmented line tool and the "Fit Spline" function. Superplots for ciliary length and swimming speed were created by measuring the mean of 3 biological replicates [99].

Genome Changes Unique to CC-5325
High confidence, unique, nonsynonymous genetic difference in coding regions of 40 genes in CC-5325 as compared to other commonly used background strains were identified using the published data in Li et al., 2016. To assess the functions of these genes, we compiled the MapMan and GO functional annotations as well as the phytozome descriptions and deflines. Putative transcription factors were identified using the Plant Transcription Factor Database (PTFDB) [100].