Methods S1
Additional details on the creation and validation of the tagged C. albicans collection.
For the mutagenesis, we used an EZ-Tn5 (Epicentre) transposon and modified it to contain i) the Gateway compatibility cassette for the TagModule transfer, ii) a kanamycin resistance marker for selection of insertion events in E. coli, and iii) a UAU1 selection cassette [1], which allows selection for integration into the C. albicans genome and can be used to generate heterozygous (Arg+) as well as homozygous disruption mutants (Arg+, Ura+). The final step to produce uniquely tagged transposons was to transfer the TagModules into the modified Tn5 vector. Because the Gateway reaction is highly efficient and specific ([2]; data not shown), TagModule transfer to the modified Tn5 transposon was performed in pools, resulting in pools of tagged transposons.
We used the pools of tagged transposons to mutagenize a commercial genomic library (Open Biosystems; [3]) in vitro. However, we found that we quickly saturated this library as few insertions in new genes were recovered after additional rounds of mutagenesis and sequencing. To improve genome coverage, we chose to use multiple, alternatively cloned genomic libraries, which were created by digesting C. albicans genomic DNA with different combinations of restriction enzymes. By using multiple libraries (thereby reducing cloning biases), we were able to improve coverage (Figure S1).
To transform the desired transposon insertion events into C. albicans to generate heterozygous alleles, the genomic fragment containing the transposon insertion was excised from the library. This flanking genomic sequence was then used for homologous recombination of the transposon insertion into the C. albicans genome. Because the average library insert size ranged from 2-8kb, we estimate that these homologous regions were generally well in excess of 60 bp, which typically yield transformation efficiencies of 97%+ in S. cerevisiae (A. Chu, personal communication). We performed a PCR-based test for integration using primers flanking regions from within the transposon to 611-726 bp outside the transposon insertion. Tests of two independent insertion events of ERG11 showed a PCR product indicating integration for 15/15 and 13/15 colonies picked, respectively; we also tested integration of ERB1 (14/15), and CDC37 insertions (22/24). We are confident that these numbers represent the lower bounds for correct integration as we used a relatively crude colony PCR-based test. Because this PCR-based method tests only for homologous integration, it is possible that there may be additional ectopic integration of the transposon-containing genomic fragment. However, given the long homologous flanking regions contained within these fragments to mediate recombination, we expect that ectopic integration would be a rare event [4].
So that there would be no overlap of tagged strains when pooled, we sorted our 21468 insertion events to maximize the number of unique genes associated with unique TagModules. We found, however, that we had fewer unique TagModules than genes available; only 3838 unique TagModules were represented across the 4827 genes. Accordingly, we asked if using one tag of a TagModule (effectively doubling the number of usable tags) would provide sufficient discrimination of individual strains in a pool. This was supported by the previous observation that the performance of both the uptag and downtag in a TagModule was robust, quantitative, and correlated [5]. Under this approach, we created two pools, termed “pool 1” and “pool 2”. Each pool has a unique set of transposon insertions but contains an overlapping subset of TagModules. Therefore, we amplify only uptags from one pool, and only downtags from the second pool. For pooled growth assays, each pool is screened and tags amplified separately prior to combining the amplicons for microarray hybridization (Figure S2A). Using this one-tag approach, we were able to select 4401 uniquely tagged genes (representing 4388 unique genes) for transformation into C. albicans (Table S1, Table S2).
To validate that this one-tag approach was reproducible regardless of whether the uptag or the downtag was used to represent a strain, we performed a “tag swap” experiment (Figure S2B). We assigned “pool 1” unique uptags, and “pool 2” unique downtags, amplifying the tags separately but combining the products prior to hybridization to an array. We compared this to the hybridization of the reverse combination and found that strains with above 3X background intensity were significantly correlated (R= 0.64, p<10-16). As reported in the main text, biological replicates of this pool grown under twenty generations of growth in a pooled assay were highly correlated (Figure 2B; R= 0.98, p<10-16). These results indicate that the hybridization performance of a strain is similar regardless of whether it was represented by its uptag or its downtag and that little information on tag abundance is lost by using only one tag. For the remainder of this study, we report data on one tag per strain.
To validate that the tags were successfully incorporated into each mutant strain following transformation into C. albicans, we tested the hybridization performance of our tagged mutants by pooling them in 384-strain batches, amplifying the tags, and hybridizing them to an array. Of the 4401 attempted transformations, we recovered colonies for 4239 of these, based on our qualification that a strain’s tag must have signal intensity above 3X median background intensity (Table S2). As reported in the main text, when we pooled all of the 4239 strains, amplified their tags, and hybridized them to an array, we detected 3633 (85%) with tag intensity above 3X median background, with 619 strains falling below background (Figure 2C).
Finally, to validate that our strains did not have significant cross-reactivity with other features on the array (due to cross contamination, sequencing errors, or errors in sample tracking), we examined the signal intensities of all unused tags on the array. The vast majority, 12292 of 12686 (97%) fell at or below 3X background (Figure S3). Of the 394 above 3X background, 166 (1.3%) corresponded to previously repaired tags, which have significant sequence similarity to tags in use [6]. 116 (0.9%) corresponded to tags with no significant sequence similarity to the tags used to create the TagModules, and 112 (0.9%) corresponded to TagModule tags that were included in the initial pool of uniquely tagged transposons, but were not included in the subsequent mutant collection. This last number may represent a low level of error in sample tracking (e.g., an incorrect tag being assigned to a strain, or cross contamination in the collection); nonetheless, the majority of these have significantly lower signal intensity relative to the expected TagModules in our pool (one-tailed t-test, p<1.7x10-12).
Pooled growth on solid SLAD media
Solid SLAD media was prepared as follows: One liter of media contained 20 g electrophoresis-grade agarose, 1.7 g yeast nitrogen base (without ammonium sulfate and without amino acids), and 20 g glucose. Following autoclaving, filter-sterilized ammonium sulfate, histidine, and uridine were added to a final concentration of 50 mM, 10 mM, and 12.5 mM, respectively. 10 mL of media was then aliquotted to 85 mm Petri dishes. The two C. albicans pools were plated independently, in duplicate, for a total of ~500,000 colonies over 15 plates. Plates were sealed with parafilm and incubated at 37°C for six days, at which point samples of agarose were examined microscopically to verify cell invasion. 3 mL of water were then added to each plate, gently rubbed with a glove, and this non-invaded portion (“supernatant”) was collected. The plates were then washed under running tap water. The remaining agarose, which contained the invaded cells, was transferred to a beaker (100 g/beaker) and 200 mL of Qiagen QG solubilization buffer added. The agarose was melted for 90 min at 60°C, after which the cells were pelleted at 10,000 RCF in a Sorvall Centrifuge (“pellet”). Samples of the supernatant and pellet cells were processed and hybridized as described [7]. Log2 ratios (supernatant/pellet) were calculated from the mean-normalized data for each replicate and averaged.
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