Atomic Force Microscopy Demonstrates That Disulfide Bridges Are

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Atomic Force Microscopy Demonstrates That Disulfide Bridges Are Required for Clustering of the Yeast Cell Wall Integrity Sensor Wsc1 Vincent Dupres,† J€urgen J. Heinisch,*,‡ and Yves F. Dufr^ene*,† †

Institute of Condensed Matter and Nanosciences, Universite Catholique de Louvain, Place Croix du Sud 1, B-1348 Louvain-la-Neuve, Belgium ‡ Fachbereich Biologie/Chemie, Universit€at Osnabr€uck, AG Genetik, D-49076 Osnabr€uck, Germany ABSTRACT: In yeasts, cell surface stresses are detected by a family of plasma membrane sensors. Among these, Wsc1 contains an extracellular cysteine-rich domain (CRD), which mediates sensor clustering and is believed to anchor the sensor in the cell wall. Although the formation of Wsc1 clusters and their interaction with the intracellular pathway components are important for proper stress signaling, the molecular mechanisms underlying clustering remain poorly understood. Here, we used the combination of single-molecule atomic force microscopy (AFM) with genetic manipulations to demonstrate that Wsc1 clustering involves disulfide bridges of the CRD. Using AFM tips carrying nitrilotriacetate groups, we mapped the distribution of individual His-tagged sensors on living yeast cells. While Wsc1 formed nanoscale clusters on native cells, clustering was no longer observed after treatment with the reducing agent dithiothreitol (DTT), indicating that intra- or intermolecular disulfide bridges are required for clustering. Moreover, DTT treatment resulted in a significant increase in cell surface roughness, suggesting that disulfide bridges between other cell-wall proteins are crucial for proper cell surface topology. The remarkable sensor properties unravelled here may well apply to other sensors and receptors with cysteine-rich domains throughout biology. Our combined method of AFM with genetic manipulations offers great prospects to explore the mechanisms underlying the clustering of cell surface proteins.

’ INTRODUCTION Fungal cell walls serve as a first barrier to adverse environmental conditions and ensure both cell shape and integrity, thus constituting a primary target for antifungal drugs.1 Among fungi, the Bakers yeast Saccharomyces cerevisiae is recognized as a model eukaryote, owing to its well-established molecular genetics. The composition of the S. cerevisiae cell wall and the signal transduction mechanisms ensuring its integrity have been extensively studied. In brief, β-1,6- and β-1,3-glucan chains form an elastic cage of carbohydrates, with minor amounts of chitin adding to its strength. Highly mannosylated cell wall proteins (CWPs) are also incorporated and basically form an outer layer linked to the polysaccharides (see ref 2 for further details). The CWPs include enzymes involved in cell wall remodelling, such as glucanases and chitinases, which are necessary for cell growth and proliferation, as well as a number of structural proteins. The latter are frequently linked to each other through disulfide bridges between cysteine residues of their peptide chains. Consequently, destroying these bridges by treatment with the reducing agent dithiothreitol (DTT) results in the release of CWPs into the medium.3,4 DTT treatment also affects the cell wall remodelling capacity.5 CWPs released from the cell wall most likely encompass proteins of the outer, electron-dense layer of the yeast cell surface, which are only loosely attached to the underlying glucan network. As expected, none of the proteins released upon DTT treatment contains a transmembrane domain (TMD), which would anchor it within the underlying plasma membrane. r 2011 American Chemical Society

However, membrane-spanning proteins also contain cysteine residues capable of forming disulfide bridges that may well be of functional relevance. Prominent examples of this type of protein are the cell wall integrity (CWI) sensors of the Wsc type, which contain a cysteine-rich domain (CRD) with eight conserved cysteines (reviewed in refs 6 and 7). Phenotypic analyses of mutants with cysteine to alanine substitutions in the CRD demonstrated that the cysteines are crucial for sensor function in vivo.8 The CRD is also frequently referred to as a “Wsc domain” and can be found in proteins in all biological systems, from viral, bacterial, fungal, and mammalian origin.9 However, neither the function nor the structure of this domain have yet been elucidated. Because the domain is also found in two fungal exoglucanases, it has been proposed to bind to cell wall polysaccharides in yeast and, thus, provide a second anchor besides their transmembrane domains for the mechanosensors.10 We recently used single-molecule atomic force microscopy (AFM)11 13 to unravel the nanomechanics14 and clustering8 of Wsc1 sensors in living S. cerevisiae cells. Clustering of Wsc1 was strongly enhanced in deionized water or at an elevated temperature, indicating its relevance in proper stress response. However, the molecular mechanisms underlying Wsc1 clustering remain poorly understood. Whether the formation of disulfide bridges within the CRD domain of Wsc1 plays a role in clustering is the Received: September 20, 2011 Revised: November 7, 2011 Published: November 22, 2011 15129

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Langmuir question we address here. To this end, we localized individual wild-type sensors on S. cerevisiae cells, either in native conditions or after treatment with DTT. We found that DTT disrupts Wsc1 clustering, indicating that intra- or intermolecular disulfide bridges are required for clustering.

’ MATERIALS AND METHODS Media and Genetic Manipulations. Yeast cell culture and genetic techniques followed standard procedures.15 Rich medium (YEPD) contained 2% glucose, 1% yeast extract, and 2% Bacto Peptone (Difco Laboratories, Inc., Detroit, MI). Synthetic media, with the omission of single bases and amino acids as required, were used for selection of plasmid maintenance. To obtain solid media, 1.5% (w/v) agarose was added prior to sterilization. Strains and Plasmids. Strain HOD48-1D (MATa wsc1::KlURA3 ura3-52 leu2-3,112 his3-11,15) and plasmid pBH01, employed in this study, have been described previously.16 In short, HOD48-1D carries a substitution of the complete WSC1 open reading frame by the URA3 marker cassette from Kluyveromyces lactis.17 pBH01 is a yeast CEN/ARS plasmid with LEU2 as a selection marker, which carries a fusion gene encoding a modified Wsc1. The modified sensor penetrates the cell wall because of an elongation by the serine/threonine-rich (STR) region of Mid2 and can be detected by a modified AFM tip because of an additional extracellular, N-terminal 8His tag.16 For AFM studies, transformants were cultured in leucine-free synthetic media as follows. Two or three colonies from the solid medium plate used as inoculum were transferred into culture medium. Cells were agitated overnight at 30 °C, grown up to the late logarithmic phase, and harvested by centrifugation. They were washed 3 times with sodium acetate buffer (sodium acetate at pH 4.75) and resuspended in 10 mL of buffer to a concentration of ∼106 cells/mL. For DTT experiments, 50 μL of a DTT solution (Sigma, 5 or 25 mM) was injected into the AFM liquid cell, giving a final concentration of 1 and 5 mM DTT, respectively. Determination of CWI Pathway Activation. The dual phosphorylation of mitogen-activated protein kinase (MAPK) Mpk1/Slt2 was employed as an indirect measure of CWI pathway activation. For this purpose, the immunological detection method described by ref 14 was modified as follows: To extract proteins, exponentially growing cells corresponding to 1 mL of cells with an OD600 of approximately 2 were collected by centrifugation, resuspended in 500 μL of 1.85 M NaOH containing 2% β-mercaptoethanol, and incubated on ice for 10 min. Trichloroacetic acid was added to a final concentration of 13% and further incubated for an additional 10 min on ice. Insoluble material was collected by centrifugation, mixed with 40 μL of sodium dodecyl sulfate (SDS) sample buffer and boiled for 5 min before separation in a 10% SDS polyacrylamide gel. After transfer to a nitrocellulose membrane (Whatman GmbH, Dassel, Germany), proteins were detected with one of two primary antisera. For detection of the total amount of Mpk1, polyclonal goat antiserum yC-20 from Santa Cruz Biotechnology (Santa Cruz, CA) was employed. For detection in the Odyssey imaging system, IRDye 800Dx conjugated affinity purified anti-goat IgC was purchased from Rockland (Gilbertville, PA). Dually phosphorylated Mpk1 was detected with phospho-specific antiserum p44/42 MAPK (Erk1/2, Thr202/Tyr204) raised in rabbits by cell signaling technology (New England Biolabs GmbH). Bound primary antibodies were visualized again in the Odyssey imaging system using IRDye 700Dx conjugated affinity purified anti-rabbit IgC from Rockland (Gilbertville, PA). Blots were visualized and quantified in the Odyssey infrared imaging system (Li-Cor, Lincoln, NE), and the software was provided by the supplier (Odyssey 2.1). AFM. AFM measurements were performed in buffered solutions (sodium acetate + 0.1 M sucrose at pH 4.75), using Nanoscope IV Multimode AFM (Veeco Metrology Group, Santa Barbara, CA) and

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gold-coated cantilevers (OMCL-TR4, Olympus Ltd., Tokyo, Japan). Cells were immobilized by mechanical trapping into porous polycarbonate membranes with 3 μm pore size (Millipore). The pore diameter was modified to match the cell diameter by etching overnight in 10 mL of 5 M NaOH (Sigma) at room temperature.18 Membranes were then rinsed several times in Milli-Q water. After filtering a concentrated cell suspension, the filter was gently rinsed with buffer, carefully cut (1  1 cm), and attached to a steel sample puck (Veeco Metrology Group) using a small piece of double-face adhesive tape, and the mounted sample was transferred into the AFM liquid cell while avoiding dewetting. We first imaged the topography of single yeast cells by engaging a noncoated tip and scanning the sample in contact mode. To detect His-tagged sensors by force spectroscopy, the tips were chemically modified with Ni2+ nitrilotriacetic acid (NTA) groups as follows. Gold-coated cantilevers were cleaned for 15 min by ultraviolet (UV) and ozone treatment, rinsed with ethanol, dried with a gentle nitrogen flow, immersed overnight in ethanol containing 0.05 mM NTA-terminated alkanethiols (ProChimia, Poland), rinsed with ethanol, further immersed in a 40 mM aqueous solution of NiSO4 (pH 7.2) for 1 h, and then finally rinsed with buffer. The spring constants of the Ni2+ NTA gold cantilevers were measured using the thermal noise method (Picoforce, Veeco Metrology Group), yielding values ranging from 0.02 to 0.025 N/m. Unless otherwise specified, all force measurements were performed using a constant approach and retraction speed of 1500 nm/s and with an interaction time of 500 ms. Adhesion maps were obtained by recording 32  32 force curves on 1  1 μm areas of the cells, calculating the adhesion force values and displaying them as gray pixels.

’ RESULTS AND DISCUSSION DTT Affects the Roughness of the Yeast Cell Surface. We first imaged the surface morphology of single yeast cells in their native state. Cells were mechanically trapped in a porous polymer membrane, thus allowing cell imaging in buffer without pretreatment, such as air drying or chemical fixation. Panels a, c, and e of Figure 1 show height and deflection images of a single native yeast cell recorded in acetate buffer. Because of the large curvature of the cell, the image obtained in the height mode (Figure 1a) has fairly poor resolution, while images obtained in the deflection mode (panels c and e of Figure 1) are much more sensitive to the surface relief. However, it should be kept in mind that deflection images do not provide quantitative height measurements. High-resolution images revealed a smooth and homogeneous surface (Figure 1e), with a root-mean-square roughness on height images (Rrms) of 1.1 nm (500  500 nm areas; Figure 2). We then explored the surface morphology of yeast cells after treatment with DTT. Panels b, d, and f of Figure 1 show the same yeast cell as in panels a, c, and e of Figure 1 but after incubation for 1 h with 1 and 5 mM DTT (inset). As can be seen, the structural integrity of the cell treated with 1 mM was slightly impaired, with a small but significant increase in surface roughness (Rrms of 1.6 on 500  500 nm areas; Figure 2). This indicates that gentle treatment with DTT did not cause substantial structural modifications. In contrast, treatment with 5 mM DTT led to a much stronger increase in surface roughness (Rrms = 2.6 nm; inset of Figure 1f and Figure 2), suggesting that this higher concentration affects the network of mannoproteins at the outer cell surface. This interpretation is consistent with previous observations, showing that some CWPs are released into the medium upon treatment with DTT.3 Besides protein release, one may also consider that a portion of the remaining 15130

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Figure 3. DTT elicits the CWI stress response only after prolonged treatment or at higher concentrations. Phosphorylation of the MAPK Mpk1 was determined by immunological detection with antisera raised against the dually phosphorylated kinase (Mpk1-P; upper row) or against the entire MAPK (Mpk1; lower row). Relative amounts of phosphorylated Mpk1-P calculated versus total Mpk1 serve as an indirect measure of CWI pathway activation. Maximal activation after treatment with 5 mM DTT for 1 h was set to 100%, and relative amounts of Mpk1-P were normalized to that value (percentage of maximal activation).

Figure 1. AFM topographic imaging reveals that the yeast surface structure is not substantially altered by treatment with 1 mM DTT. AFM height (a and b; z range = 2 μm) and deflection (c f) images of the surface of S. cerevisiae recorded in buffer before (a, c, and e) and after (b, d, and f) injection of 1 mM DTT for 1 h. For comparison, the inset image in panel f was obtained after injection of 5 mM DTT.

Figure 2. Higher amounts of DTT increase the yeast cell wall roughness. Statistical analysis of topographic images obtained from S. cerevisiae cells prior to and after treatment with 1 and 5 mM DTT for 1 h. Variations of Rrms were constructed from power spectral density analysis of the height images as a function of the length scale. Each data point represents the mean ( standard error of the mean (SEM) of five images obtained on five different cells.

proteins change their conformations because of breakage of disulfide bridges, thus also affecting surface roughness and

electrostatic charges. Cell surface charges could then enhance electrostatic interactions between the tip and sample, therefore contributing to the observed differences between treated and nontreated cells. Moderate Treatment with DTT Does Not Induce CWI Signaling. To determine the effect of DTT on the CWI signaling pathway in cultured yeast cells, we assessed the dual phosphorylation of the downstream MAPK Mpk1/Slt2 as a measure of pathway activation. Environmental stress, such as heat, low medium osmolarity, or the addition of antifungals, such as the echinocandin caspofungin, has been shown to elicit a massive phosphorylation of the MAPK19 (reviewed in ref 20). As evident from the data in Figure 3, treatment with 1 mM DTT for 30 min did not cause sufficient stress on the yeast cells to rapidly induce the CWI response. Rather, Mpk1/Slt2 phosphorylation stayed at the baseline level of the control culture without DTT treatment. After 60 min of incubation with 1 mM DTT, an intermediate phosphorylation level was detected, indicating that the increase in cell surface roughness detected above triggers some CWI pathway activation. Clearly, treatment with higher DTT concentrations (5 mM) elicited an immediate and strong phosphorylation of Mpk1/Slt2, i.e., posing a significant cell surface stress. Again, this is consistent with previous observations of several CWPs being released into the medium upon DTT treatment.3,4 The coincident loss of structural integrity of the cell wall is expected to trigger mechanosensing through Wsc1 and the other homologous sensors.7 Moreover, we have previously shown that cell surface stress leads to a massive increase in the number of Wsc1 sensors and its clustering.8 Single-Molecule AFM Reveals Wsc1 Clustering on Live Cells. We next investigated the distribution of single Wsc1 sensors with the aim to determine whether it is altered upon DTT treatment. Single-molecule Wsc1 detection by AFM employed the following strategy: the use of chemically modified AFM tips, combined with a modification of the gene encoding the Wsc1 sensor, so that the protein reaches the cell surface because of an additional STR region introduced from another sensor and can be specifically picked up by the tip (Figure 4a). The site-directed NTA polyhistidine (Hisn) system has proven to be particularly well-suited for single-molecule AFM detection.21,22 The NTA Hisn binding chemistry is well-known and involves the formation of a hexagonal complex between the tetradental ligand NTA and divalent metal ions, such as Ni2+. Because NTA occupies four of the six coordination sites of Ni2+, 15131

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Figure 4. AFM detects and force probes single Wsc1 sensors on live cells. (a) His-tagged, modified Wsc1 membrane sensors were detected using AFM tips functionalized with Ni2+ NTA groups. The drawing shows a His-tagged, elongated Wsc1 sensor, with the cytoplasmic tail (CT), the transmembrane domain (TMD), the cysteine-rich domain (CRD), the STR region, and the terminal His tag (in green). cw = cell wall, and pm = plasma membrane. (b and d) Adhesion force histograms (n = 4096) and representative force curves recorded with a Ni2+ NTA tip in buffer solution (sodium acetate + 0.1 M sucrose + 40 mM NiSO4 at pH 4.75) for S. cerevisiae cells expressing His-tagged, elongated Wsc1 sensors (b) prior to and (d) after treatment with 1 mM DTT for 1 h. (c and e) Representative force extension curves obtained upon stretching single Wsc1 (c) prior to and (e) after treatment with 1 mM DTT for 1 h. The curves displayed a linear region, where force is directly proportional to extension, thus characteristic of a Hookean spring. Using the slope of the linear portion of the raw deflection versus piezo displacement curves, we found that the spring constant ks of Wsc1 did not change with DTT treatment and was in the range of 4.4 4.6 pN nm 1.

the two remaining sites are accessible to other Lewis bases, e.g., the histidines of tagged proteins. In the AFM context, the method can be readily applied to live cells, e.g., for fishing recombinant histidine-tagged proteins expressed on the cell surface using AFM tips coated with NTA groups. Owing to this novel procedure, it is possible to force probe by AFM not only natural surface proteins but also those usually located well beneath the surface within the cell wall. His-tagged sensors were detected by scanning the cell surface with an AFM tip bearing Ni2+ NTA groups. As shown in Figure 4b, a substantial fraction (10%) of the force curves recorded across the cell surface displayed single adhesion force peaks, with the remaining curves exhibiting no adhesion. The corresponding adhesion force histogram (Figure 4b) displayed force values that were typically in the 100 200 pN range. In light of previous studies,14,21 we attribute these forces to the rupture of a single NTA His bond and, thus, to the detection of a single His-tagged sensor. The few adhesion events detected in the 200 400 pN range are likely to reflect the detection of two sensors. The specificity of such analyses was confirmed by showing a dramatic reduction of adhesion events in the absence of Ni2+ or on related yeast strains lacking the sensors. Consistent with earlier work,14 pulling on sensors produced specific force distance curves, showing a first regime corresponding to the straightening of the extracellular chain at very low force, followed by a characteristic linear region, where force is directly proportional to extension, thus typical of a Hookean spring (Figure 4c). As shown in Figure 5a, adhesion maps recorded over 1  1 μm areas revealed the distribution of the sensors at the cell surface. There are two important comments associated with these recognition maps: first, every bright pixel in a map represents the detection of single sensors mostly, sometimes two, consistent with the adhesion histogram (Figure 4b), and second, the apparent

surface density of the sensors, 92 ( 9 sensors/μm2 (n = 9; Figure 5e), represents a lower estimate because it is possible that more sensors are exposed but not detected, owing to experimental limitations [lateral resolution of ∼30 nm (pixel size), His tag not properly oriented, etc.]. Importantly, we found that many sensor molecules were assembled into clusters. Dimensions of the clusters were in perfect agreement with those previously found under the same conditions.8 In that earlier work, we also showed that both the total amount of wild-type Wsc1 sensors and their frequency of clustering increased upon environmental stress, i.e., application of either heat or low osmolarity. This finding was fully consistent with the notion that stress conditions activate the CWI pathway and, thus, indicate that clustering is a stress-responsive process that is intimately connected to signaling. Wsc1 Clustering Is Disrupted by DTT. To address the question of whether Wsc1 clustering is sensitive to reducing conditions, we explored the cell surface distribution of Wsc1 after treatment with 1 mM DTT. Panels d and e of Figure 4 show that DTT did not alter the overall density of sensors at the surface or their spring behavior (sensor spring constant k ≈ 4.5 pN nm 1). In contrast, panels b e of Figure 5 show that clustering was strongly reduced on DTT-treated cells, while the overall density of sensors at the surface remained constant. This effect was stable over time because adhesion maps recorded several hours after DTT injection still showed a lack of clustering (panels c and d of Figure 5). These findings are complementary to the results of earlier experiments, in which we mapped the distribution of mutants within the CRD domain of Wsc1.8 There, specific cysteines within this domain had been replaced by alanine residues, leading to a lack of clustering and a simultaneous loss of in vivo signaling function. Nevertheless, such mutants displayed adhesion frequencies, adhesion values, and spring behaviors similar to those 15132

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turnover by delivery to and removal from the surface, with the latter caused by endocytosis.23 Clustering could thus result from the localized delivery of sensors from intracellular vesicles but would presumably still require some lateral movement within the fluid membrane, mediated by their transmembrane domains. To which degree this movement is influenced by the long extracellular regions of the sensors has not yet been investigated. However, because we demonstrated earlier that the elongation by the additional STR and the His tag does not contribute to the sensor mechanics,14 we believe that these modifications would alter the kinetics at the cell surface only to a minor extent.

’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected] (J.J.H.); yves.dufrene@ uclouvain.be (Y.F.D.).

Figure 5. Single-molecule AFM demonstrates that Wsc1 clustering is disrupted by DTT. (a d) Representative adhesion force maps obtained by scanning 1  1 μm areas with a Ni2+ NTA tip in buffer solution (a) before and (b d) after injection of 1 mM DTT. Bright pixels represent the detection of single sensors; nanoscale clusters are highlighted by dotted red lines. We define a cluster as a group of sensors containing at least 10 bright pixels in either direct contact with each other or separated by no more than one dark pixel. The first map upon DTT treatment was recorded (b) 30 min after injection, while the following two were obtained after (c) 1 h and (d) 4 h. The data shown are representative of results obtained on four different cells using four different tips. (e) Surface density histograms showing the number of sensors per micrometer squared measured for Wsc1 in buffer (left) before (n = 5 maps containing 1024 data points each) and (right) after DTT treatment (n = 9 maps containing 1024 data points each). Darker and lighter colors represent the surface density of clustered and isolated sensors, respectively.

of the wild-type protein, indicating that sensor mechanical properties are exclusively determined by the STR region. The perturbation of clustering by the addition of DTT as observed here indicates that the cysteines within the CRD domain are indeed involved in mediating protein interactions, presumably within neighboring sensors but possibly as well between sensors and other CWPs. We propose that the cysteines form intramolecular disulfide bridges, which ensure a conformation of the protein surface capable of such interactions. Mutations in single cysteines or disruption of the disulfide bridges by DTT treatment would then alter the conformation and thereby disrupt the interactions. This mechanism is consistent with the fact that CRD domains heterologously produced in Escherichia coli are not able to interact with yeast cell wall material, probably because they do not pass the secretory pathway, in which the disulfide bridges are formed in S. cerevisiae and other eukaryotes. Another mechanism that cannot be ruled out yet by our data is that intermolecular disulfide bridges are formed between different Wsc1 sensor molecules or with other CWPs, which would also be disrupted by the cysteine mutations or DTT treatment. A pertinent question remaining to be addressed is the movement of the sensors at the cell surface. Sensors are subject to a rapid

’ ACKNOWLEDGMENT Work in the Yves F. Dufr^ene team was supported by the National Foundation for Scientific Research (FNRS), the Foundation for Training in Industrial and Agricultural Research (FRIA), the Universite Catholique de Louvain (Fonds Speciaux de Recherche), the Federal Office for Scientific, Technical, and Cultural Affairs (Interuniversity Poles of Attraction Programme), and the Research Department of the Communaute Franc-aise de Belgique (Concerted Research Action). Yves F. Dufr^ene is Senior Research Associate at the FNRS. Work in the J€urgen J. Heinisch team was funded by a grant from the Deutsche Forschungsgemeinschaft to J€urgen J. Heinisch within the framework of the SFB944. We thank Bernadette Sander-Turgot for excellent technical assistance in the experiments on the immunological detection of the Mpk1 phosphorylation state. ’ REFERENCES (1) Heinisch, J. J. Biochim. Biophys. Acta, Proteins Proteomics 2005, 1754, 171–182. (2) Klis, F. M.; Brul, S.; De Groot, P. W. J. Yeast 2010, 27, 489–493. (3) Cappellaro, C.; Mrsa, V.; Tanner, W. J. Bacteriol. 1998, 180, 5030–5037. (4) Insenser, M. R.; Hernaez, M. L.; Nombela, C.; Molina, M.; Molero, G.; Gil, C. J. Proteomics 2010, 73, 1183–1195. (5) Popolo, L.; Ragni, E.; Carotti, C.; Palomares, O.; Aardema, R.; Back, J. W.; Dekker, H. L.; de Koning, L. J.; de Jong, L.; de Koster, C. G. J. Biol. Chem. 2008, 283, 18553–18565. (6) Rodicio, R.; Heinisch, J. J. Yeast 2010, 27, 531–540. (7) Jendretzki, A.; Wittland, J.; Wilk, S.; Straede, A.; Heinisch, J. J. Eur. J. Cell Biol. 2011, 90, 740–744. (8) Heinisch, J. J.; Dupres, V.; Wilk, S.; Jendretzki, A.; Dufrene, Y. F. PloS One 2010, 5, 9. (9) Ponting, C. P.; Hofmann, K.; Bork, P. Curr. Biol. 1999, 9, R585–R588. (10) Heinisch, J. J.; Dufrene, Y. F. Integr. Biol. 2010, 2, 408–415. (11) Muller, D. J.; Helenius, J.; Alsteens, D.; Dufrene, Y. F. Nat. Chem. Biol. 2009, 5, 383–390. (12) Muller, D. J.; Dufrene, Y. F. Curr. Biol. 2011, 21, R212–R216. (13) Muller, D. J.; Dufrene, Y. F. Trends Cell Biol. 2011, 21, 461–469. (14) Dupres, V.; Alsteens, D.; Wilk, S.; Hansen, B.; Heinisch, J. J.; Dufrene, Y. F. Nat. Chem. Biol. 2009, 5, 857–862. (15) Sherman, F.; Fink, G. R.; Hicks, J. B. Laboratory Course Manual for Methods in Yeast Genetics; Cold Spring Harbor Laboratory: Cold Spring Harbor, NY, 1986. 15133

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