We generated S. cerevisiae Cdc12-conGFP (AGY169; DHD5 yeast with AGB467: pRS416-ScCDC12-conGFP3) based on rationale from previous work (Vrabioiu and Mitchison, 2006, 2007; DeMay et al., 2011a,b). pRS416-ScCDC12-conGFP3 was generated by removing nucleotides encoding 3 amino acids of the N-terminal alpha-helix of GFP and 4 amino acids of the C-terminal region of Cdc12. AGB455 (pScCDC12) was generated by amplifying ScCDC12 from S. cerevisiae genomic DNA using the oligonucleotides AGO1181 and AGO1182. The PCR product was ligated into the pRS416 plasmid (AGB441) and verified by test digestions using PspXI and SlaI and sequencing (Dartmouth College Core Facility, Hanover, NH). Next, the plasmid AGB459 (pScCdc12-GFP) was generated by first amplifying the GFP insert from the plasmid AGB005 using the oligonucleotides AGO1187 and AGO1188 and ligating the product into the digested AGB455 plasmid. Again this plasmid was verified with test digestions and sequencing (Dartmouth College Core Facility, Hanover, NH). Constraining of GFP to Cdc12 was completed using PCR to amplify the entire AGB459 using the oligonucleotides AGO1203 and AGO1204 which contain homology over the CDC12-GFP linker and this amplification removed nucleotides encoding 4 amino acids from the N-terminus of GFP and 3 amino acids from the C-terminus of Cdc12. AGB467 (pRS416-ScCDC12-conGFP) was verified by sequencing (Dartmouth College Core Facility, Hanover, NH). Once verified, AGB467 was transformed into wild-type lab strain (AGY000), and the following yeast deletion strains: bnr1Δ, bni1Δ, bud4Δ, cla4Δ, gin4Δ, rts1Δ, and shs1Δ (kindly provided by Erfei Bi, AGY028-AGY035 to generate AGY066-AGY073). Doug Kellogg generously supplied us with shs1 phosphomutant strains (Egelhofer et al., 2008) and the plasmid AGB467 was transformed into these strains as well to generate AGY317-AGY320. To generate strains to measure septin intensity over the cell cycle the plasmid AGB553 (E1915 YIplac128-GFP-ScCDC3:LEU a gift from Erfei Bi) was integrated into the same strains as the AGB467 plasmid generating the strains AGY131-AGY137, and AGY323-AGY327. All strain, plasmid, and oligonucleotide information is present in Tables 1–3.

For imaging, S. cerevisiae cells were grown overnight in appropriate media with proper selection for the specific plasmids, collected by centrifugation, and resuspended in 2x low fluorescence minimal media (LFMM). Cells were mounted on gel pads of 1.4% agarose and LFMM on glass depression slide, covered with a coverslip (no. 1.5), sealed with Valap, and imaged.

Time-lapse recordings for estimating septin intensity of mutant strains through the cell cycle were acquired with a Nikon Eclipse Ti-E inverted wide-field microscope equipped with a 60x (1.4 N.A.) plan-apochromat oil objective and a Andor Zyla 4.2 sCMOS camera. A Chroma DAPI/FITC/TRITC/Cy5 quad filter set was used for fluorescent imaging of GFP. The fluorescent light source was a Spectra LED lamphead and images were acquired with 12% laser power, 100 ms exposure, and 90 s time intervals.

Using FIJI, individual cells were cropped from the timelapse recordings and the mean background-subtracted septin intensity at the mother-bud neck were determined for every time point for a given cell using MATLAB. These septin intensity values were saved and plotted in an Excel file. For every cell, a time zero (t0) was manually determined. This t0 represented the maximum septin intensity value close (not more than 4–5 time points) to the visible ring split. In most cases the quantitatively determined t0 was not more than two time points (3 min) away from the visible split. This quantitative approach was taken to account for different visible split times depending on how a given image was automatically contrasted by FIJI. t0 therefore represented the quantitative onset of ring splitting. Since the primary focus of this study was the septin concentration change during ring splitting, all following analyses were limited to 24 min before and after t0. Percent change over the entire 24 min was determined and reported in Table 4. Since the majority of the strains analyzed showed biphasic septin disassembly rates over the course of ring splitting, the disassembly rate was determined for both phases of disassembly (phase 1 and phase 2). Phase 1 and phase 2 were distinguished by the different slopes of the biphasic intensity curves. If a strain did not show biphasic disassembly, then it is reported as monophasic and the intensity from t = 0 to t = 24 min after splitting; Phase 2 is the rate of disassembly from the last time point of Phase 1 to t = 24. After disassembly rates were determined, the fold change compared to wild-type was calculated.

All polarization fluorescence images (except the shs1-phosphomutant strains) were acquired using the MF-PolScope (Abrahamsson et al., 2015). The MF-PolScope was set up on an inverted Olympus IX-83 microscope body, equipped with an Olympus 60x (1.3 N.A.) silicon-oil immersion objective, and the LC-compensator for polarization-controlled excitation light, with a green 515/30 nm emission filter (Semrock) placed before an EMCCD camera (Andor iXon-888). The fluorescence light source used was a blue LED module of an X-cite XLED1 (Lumen Dynamics). Images were acquired through Micro-manager (www.micro-manager.org) using the OpenPolScope software (www.openpolscope.org).

Polarized fluorescence measurements of the shs1-phosphomutant strains and bud4Δ splitting events were acquired with the Zeiss Axio Imager.M1 wide-field microscope with five linear polarized filters (Chroma 21033a) in a filter wheel (Ludl Electronic Products, cat. no. 99A075) and UV blocker 420 nm LP emission filter 32 nm (Chroma, cat. no. E420LPv2-32). The fluorescent light source used was an EXFO X-Cite 120 lamp. Images were acquired through Micro-manager (www.micro-manager.org) using the OpenPolScope software (www.openpolscope.org).

Once acquired, all polarization images were analyzed using the OpenPolScope software. Analysis involved internal background corrections of every cell, performed by selecting a ROI in the cytoplasm, and a ROI in the septin hourglass structure and then processed for anisotropy. The anisotropy formula calculates Imax-IminImax+Imin in each pixel, based on the bleach corrected fluorescence intensity values. Circular statistics were used to calculate the ensemble orientation and anisotropy for each given septin structure. To do this, a 6 × 6 pixel ROI was selected in the center of the septin structure so as to exclude edges of the septin structure with isotropic measures from GFP β-barrel orientation along the curve of the septin ring. The pixel-by-pixel orientation, anisotropy, and intensity for each septin structure was then exported, and used to calculate the ensemble orientation weighted by anisotropy and intensity of each ring, and of the entire population of cells. In addition we also calculated the population variance (PopVar=1-(Vector Average Anisotropy*IntensityScalar Anisotropy*Intensity)). A detailed description of how to use the OpenPolScope software for polarization imaging and analysis is present in Current Protocols in Cell Biology (McQuilken et al., 2015).

Previous data from our lab and others have shown that there is a 90° reorientation of the organization within the septin hourglass structure over the course of septin ring splitting at cytokinesis in S. cerevisiae cells (Vrabioiu and Mitchison, 2006; DeMay et al., 2011a,b). We had hypothesized in our work that one explanation of the apparent reorganization is that there are two populations of septins in the hourglass and this could be seen by Ong et al. (2014) at least at the onset of cytokinesis. Upon ring splitting, one of these sub-populations is thought to leave the bud neck and another population assembles and/or is retained perpendicular to the original population. What drives the disassembly and reassembly process of septins within such a small region of the cortex in a short window of the cell cycle?

We hypothesized that whether a given septin filament is released or stays localized at the neck could be explained by the composition of septin heteromeric rods, specifically the terminal subunit, which is occupied by either Cdc11 and/or Shs1. If Cdc11 or Shs1 specified which complexes were retained or exited from the neck, we predicted that Cdc11 and Shs1 would display different rates of change in concentration at splitting. To assess this, we monitored the abundance of 3 different septins at the mother-bud neck over the course of septin ring splitting. We used widefield fluorescence microscopy to image GFP-Cdc3, Shs1-GFP (in two different strain backgrounds), and Cdc11-GFP every 90 s to ensure sufficient time resolution for assessing intensity differences and kinetics (see Materials and Methods). Using two different Shs1-GFP strains we observed the accumulation of Shs1 in the septin hourglass is only ~50% of the level of either Cdc3 or Cdc11 (Figure 1A). Interestingly, we also observed that the decrease of all three septins over the course of ring splitting occurred in a biphasic manner. The first phase occurred within the first ~10 min after ring splitting. This first phase of septins leaving the bud neck is consistent with the timing of the isotropic “transition” period observed previously (DeMay et al., 2011a). Time-lapse imaging indicated that all septin proteins decrease in abundance with comparable rates during ring splitting and decrease by ~50% in concentration over the first phase of disassembly (phase 1), regardless of their starting abundance (Figure 1A, Table 4, and consistent with what was seen for Cdc3 in Dobbelaere et al., 2003; Wloka et al., 2011). When we measure the difference in disassembly rates over the first phase of disassembly between the different septins there was little difference in the rates (Table 4). Additionally, when Cdc11 and Shs1 are tagged in the same cell with different fluorophores, the decrease in abundance at the neck is nearly simultaneous for each protein, further indicating that these two subunits behave similarly with regard to septin dynamics in this part of the cell cycle (discrepancy between times of −0.3 ± 0.3 min, N = 40 cells, difference not significant from 0 with p > 0.41). These data indicate that the difference between the septin populations that stay associated with the cortex and that are released into the cytoplasm is not simply the presence or absence of Cdc11 or Shs1.

We next used time-lapse imaging and polarization microscopy to assess the contribution of septin interacting proteins on the organization and behavior of septins in the splitting transition. We first tested the hypothesis that a subset of septins could be specified to disassemble or be anchored via connections with F-actin assemblies. The yeast formin Bnr1 is regulated by Shs1 indicating these two factors are functionally interacting at this window of the cell cycle (Buttery et al., 2012). We assessed how the two formins, Bni1 and Bnr1, influence septin intensity changes and reorganization over septin ring splitting (Figure 1B). In strains lacking formins we observed a decrease in the initial intensity of septins in the hourglass, suggesting the actin cytoskeleton contributes partially to the accumulation of septins at the mother bud neck prior to ring splitting. This defect was most severe in bni1Δ, where septin intensity at the hourglass was reduced by ~65% relative to the hourglass in wild-type cells. However, the rate at which the septin sub-population exited the bud-neck during ring splitting was again observed as a biphasic disassembly, was not substantially affected in any of the strains (0.7-fold faster for phase 1 and 0.5-fold slower for phase 2 in bni1Δ and 0.6-fold faster in phase 1 and no fold change difference for phase 2 bnr1Δ, Table 4) consistent with previous reports (Feng et al., 2015).

We next asked if the formins were required for the characteristic 90° reorganization. To address this, we used polarization microscopy to measure the organization of the septin hourglass and split rings in wild-type, bni1Δ, and bnr1Δ strains containing Cdc12-conGFP (see Materials and Methods, Figures 1C–E). Specifically, we used multifocus (MF-PolScope) microscopy to simultaneously capture different Z-positions along with polarized fluorescence information so as to determine if orientation was comparable at all positions in the ring at the same time (Abrahamsson et al., 2015). When the orientation is consistent between Z-slices of the top, middle, and bottom of septin assemblies, we have interpreted this to mean that the prevailing population of septins are “paired filaments” (DeMay et al., 2011a,b).

As previously published from our lab, the septin hourglass of wild-type cells expressing Cdc12-conGFP exhibit GFP dipoles with a net orientation perpendicular to the mother-bud axis, while septin rings exhibit a net orientation parallel to the mother-bud axis (DeMay et al., 2011a,b). The net orientation of GFP dipoles in bni1Δ and bnr1Δ are comparable to wild-type on average however the anisotropy is lower for both mutants suggesting a slightly less well-ordered structure (Figure 1D, Table 5). We categorized each orientation data point by the state of the ring (broken, aberrant size) but could not see any correlation between grossly misorganized rings and orientations substantially deviating from 90° (Figure 1D). There seems to be little difference between the net orientation in the top, middle, and bottom focal planes in the two formin mutants suggesting pairing or symmetric organization relative to the plasma membrane (Figures 1E,F). In addition, we also estimated the population variance, which is a measure of the difference in average dipole orientation between cells with the same type of structure, and found little difference in each strain over the entire population compared to wild-type cells (Figure 1G). These results suggest that although Bni1 and Bnr1 are required for the proper accumulation and organization of septins in the septin hourglass, they are not required for proper timing of disassembly and reorientation of septins at ring splitting (Figures 1B–E).

We next examined other septin interacting proteins beginning with Bud4, an anillin-like protein. Bud4 has been shown to associate with septins and mutants do not have two split septin rings making them a key candidate in regulation of septin orientation in this process (Wloka et al., 2011). We looked at septin intensity through time over ring splitting in the bud4Δ mutant. We observed septins leave the bud neck in a monophasic manner (2-fold faster than wild-type in phase 1, and 3.5-fold faster than wild-type in phase 2). This rapid disassembly from the bud neck is consistent with previous data that Bud4 is required for stability of split septin rings during and after cytokinesis (Figure 2A, Table 4, Wloka et al., 2011; Eluère et al., 2012; Kang et al., 2013). The ring disassembled asymmetrically with one side much faster than the other but the analysis here was averaged across the whole ring to be consistent with other measurements in the study. Similar to bnr1Δ mutants, bud4Δ mutants showed a reduced septin accumulation within the hourglass with ~70% the intensity observed in wild-type hourglasses.

Using polarization microscopy, we observed little difference in terms of septin hourglass orientation, a slight decrease in anisotropy, and comparable population variance across all focal planes compared to wild-type (Figures 2B–F, Table 5). Any variability in GFP dipole orientation observed in the bud4Δ mutant could not be explained by visible septin defects (Figure 2C). However, when we analyzed the orientation of split rings in the bud4Δ we observed a severe decrease in anisotropy, across all focal planes suggesting that Bud4 is also required for proper organization of septin split ring structures (Figures 2C–F, Table 5). This is again consistent with previous data that Bud4 is required for the stability of the septin split rings (Wloka et al., 2011; Eluère et al., 2012; Kang et al., 2013).

The isotropic transition between the two orthogonal organization states is quite transient, lasting just a few minutes, indicating that the cue to direct disassembly and/or reassembly could be a post-translational modification. Recipients of the modification could be a subpopulation of septins, a septin regulator and/or modifiers of the local membrane composition. We examined two kinases, Cla4 and Gin4, and one phosphatase, Rts1, that have been shown to contribute to septin ring behavior (Longtine et al., 2000; Mortensen et al., 2002; Dobbelaere et al., 2003; Gladfelter et al., 2004; Versele and Thorner, 2004; Roelants et al., 2015). Both gin4Δ and cla4Δ cells showed a decrease in septin accumulation in the hourglass, both exhibiting ~50% reduction in intensity relative to the wild-type hourglass (Figure 3A, green and gold lines). In addition, we observed that septins do not leave the bud neck in a biphasic manner in the gin4Δ mutant (Figure 3A). Instead, septins leave the bud neck at a constant rate that is slower in both phases compared to wild-type cells (~75% the rate of wild-type in phase 1, and ~30% slower than the rate wild-type in phase 2, Table 4). In contrast, in cla4Δ cells septins still left the bud neck in a biphasic manner similar to the rate seen in wild-type cells (0.1-fold faster than wild-type in phase 1, and 0.4-fold faster than wild-type in phase 2, Table 4). Notably, Rts1 had little effect on septin accumulation in the hourglass, and no effect on disassembly rate relative to wild-type cells (Figure 3A, Table 4). Thus, Gin4, and to a lesser extent Cla4, are required for the sharp decrease in septin abundance at the neck that accompanies ring splitting.

We next used polarization microscopy to measure the organization of the septin hourglass and split rings in cla4Δ, gin4Δ, and rts1Δ mutant strains (Figures 3B–D). In cla4Δ and rts1Δ cells the spread of orientations for the GFP-dipole in the hourglass increased (thus the population variance increased), but the net orientation remained ~90° with some decrease in anisotropy (Figures 3C,D,F, Table 5). In both cla4Δ and rts1Δ strains the split structure was not appreciably different from wildtype (Figures 3B–D). Unlike cla4Δ and rts1Δ, organization within the septin hourglass in the gin4Δ mutant was highly disorganized, while the split structure maintained its ordered, net orientation of ~0/180° (Figures 3B–D). This data indicates that Gin4 is required for ordered septin hourglass organization and efficient disassembly but not the split ring organization. As with bni1Δ, bnr1Δ, and bud4Δ, little difference between the net orientation in the top, middle, and bottom focal planes was observed, and the increase in variability in GFP-dipole orientations within the population could not be correlated with visible septin higher-order structure defects because morphologically aberrant and normal appearing rings had a similar spread of orientations (Figures 3C,E). These data are consistent with a well-appreciated role for Gin4 in establishing the organization of septins but also indicates it is involved in executing the switch-like disassembly process. However, the septins that are recruited/retained after ring splitting do not require Gin4 for normal orientation or order.

It is possible that phosphorylation on a subset of septin rod complexes could generate the two distinct pools of septins at cytokinesis that are either in a state of disassembly or stay assembled at the membrane. As Shs1 has the most evidence of phosphorylation, we examined septin organization in cells lacking Shs1 (Mortensen et al., 2002; Egelhofer et al., 2008).

We first looked at septin intensity over ring splitting in a shs1Δ mutant strain. We observed a defect in the accumulation of GFP-Cdc3 to the septin hourglass (~70% reduction compared to wild-type intensity), and similar to what we observed in gin4Δ mutant cells, a monophasic disassembly rate of septins from the bud neck slower than either observed phase in wild-type cells (0.6-fold slower than wild-type phase 1, and 0.3-fold slower than wild-type phase 2, Table 4, Figure 4A) When observed by polarization microscopy, there was a dramatic increase in the disorganization of both the hourglass and split ring structures in the shs1Δ mutant strain, consistent with what has been seen for shs1Δ mutants by cryo-EM tomography and platinum replica EM (Figures 4B–D, Bertin et al., 2012; Ong et al., 2014). Septin disorganization was reflected in a wide spread of GFP-dipole orientations and thus an increased variance within the population, and a decrease in anisotropy (Figures 4D,F, Table 5). As with the previous mutant strains we analyzed, the difference in spread of angles was not associated with visible defects in septin structures, and no obvious difference was observed in GFP-dipole net orientation between the top, middle, and bottom focal planes (Figures 4C,E).

It is clear that complete loss of Shs1 leads to gross abnormalities in the construction and organization of septin structures throughout the cell cycle making it difficult to disentangle assembly defects from a role of Shs1 in the splitting and reorganization process. We hypothesized the highly phosphorylated C-terminus of Shs1 could be contributing to septin organization. We therefore, examined cells carrying shs1ΔC, where the C-terminus of Shs1 is eliminated. shs1ΔC cells showed a similar decrease in septin accumulation in the hourglass as the shs1Δ null strains, but interestingly still showed biphasic disassembly of septins over the course of splitting (Figure 4A, Table 4). When septin organization was assessed using polarization microscopy, both hourglasses and split rings in shs1ΔC were disorganized, similar to shs1Δ (Figure 4, Table 5). Taken together these data suggest the C-terminus of Shs1 is not important for the biphasic exit of septins from the mother-but neck but is necessary for proper organization of septin higher order structures throughout the cell cycle. What aspect of the Shs1 C-terminus makes it important for septin organization but not septin disassembly?

We next analyzed the organization of septin higher-order structures in mutants of Shs1 that have altered phosphorylation sites (Egelhofer et al., 2008). Several shs1 phosphorylation defective mutants were evaluated that had been previously generated based on results from SILAC mass spectrometry experiments in the Kellogg lab. These were ps1 (Pho85 sites), ps2 (CDK sites), and ps4 (Pho85/CDK sites) (Figure 5A; Egelhofer et al., 2008). Septin disassembly rates were notably faster in the wild-type strain background of these mutant alleles, which could be due to background mutations such as in BUD4, which is consistent with the observed asymmetric splitting also observed in certain bud4Δ strains (Voth et al., 2005). The shs1 ps mutants showed strikingly distinct assembly and disassembly defects amongst each other. shs1-ps1 and shs1-ps4 had substantially diminished levels of septins in assembled hourglasses, ~50 and 80% reduction in intensity relative to wild-type, respectively (Figure 5B). In contrast, shs1-ps2 cells had similar septin levels at the hourglass compared to wild-type as gauged by GFP-Cdc3 intensity. Interestingly, none of the shs1 ps mutants had a different rate of disassembly from wild-type (Table 4). This is potentially a result of the strain background interfering with any differences in rates relative to one another; however, these data still suggest that phosphorylation may be able to tune the affinity of septins for membrane at the neck or septin-septin interactions for the assembly of the hourglass.

All the shs1 phosphorylation-deficient mutants were comparably highly disorganized and showed a wide distribution of orientations and cell-to-cell variability in both hourglass and asymmetric split ring structures (Figures 5C–F, Table 5). The impact of these mutations is comparable to a complete null mutant of SHS1. These data attest to the key importance of Shs1 and likely its regulation by phosphorylation in modulating the abundance and organization of septin higher-order structures but do not support a major role of these modifications in the splitting process.