Research reported in this publication was supported by the National Institute of General Medical Sciences of the National Institutes of Health under award number GM060221 and in part by the National Institute of General Medical Sciences of the National Institutes of Health under award number T32GM007223. R.C. was supported in part by the UNC-Charlotte Faculty Research Grants Program.

1. Yeast Strain Construction
<ol>
	<li>Begin with TVY614 (pep4&#916;::LEU2 prb1&#916;::hisG prc1&#916;::HIS3)11 as the parent strain. This strain is deficient for vacuolar proteases, which contribute to the majority of protein degradation after cell lysis, and therefore allows for a cleaner and more efficient purification.</li>
	<li>Design primers12 and integrate tandem affinity purification (TAP) tag at the C-terminus of Atg20 (SNX-BAR ORF 1) using homologous recombination. Use polymerase chain reaction (PCR) to confirm integrations (Figure 2).</li>
	<li>Perform a western blot of cell lysate against the TAP tag to confirm proper integration13. 
	NOTE: We recommend harvesting 3 OD (1 OD &#8776; 1 x 107 cells) of cells for SDS-PAGE and western blot verification. Note that integration of the TAP tag should occur prior to replacing the endogenous promoters with the GAL1 promoter to allow for easier TAP tag verification via western blot.</li>
	<li>Replace endogenous Snx4 (SNX-BAR ORF1) and Atg20 (SNX-BAR ORF 2) promoters with untagged GAL1 promoter using sequential homologous recombination and transformation steps of each individual ORF14.</li>
	<li>Use flanking primers outside of the integration sites to PCR confirm successful integrations (Figure 2). This will result in a null phenotype of the targeted SNX-BARs in the absence of galactose in the growth medium.</li>
</ol>
2. Yeast Induction and SNX-BAR Dimer Purification
NOTE: Yeast cells can be propagated on standard YPD (yeast extract, peptone, and 2% glucose) agar plates as the modifications are chromosomally integrated.
<ol>
	<li>Inoculate a large swab of cells into 50 mL of standard YP (yeast extract and peptone) medium with 2% raffinose and 0.1% glucose as carbon source in a flask at least 4x the volume of the culture and grow overnight in 30 &#176;C shaker to allow proper aeration. Expect that growth in this medium will be slower compared to standard YPD.</li>
	<li>Next morning, use the 50 mL preculture to inoculate into 1 L of standard YP medium with 2% raffinose and 0.1% glucose and grow for 4-5 h in 30 &#176;C shaker. 
	NOTE: The volume of preculture used to inoculate 1 L culture may be adjusted depending on the OD600 of the preculture. The OD600 of the 1 L culture after inoculation should be around 0.2 to allow for at least two doublings during the 4-5 h growth.
	<ol>
		<li>Use a baffled Fernbach flask for growth to allow proper aeration, a 2.8 L volume flask is sufficient. Less aeration may result in slower growth and smaller cell pellet upon harvesting.</li>
	</ol>
	</li>
	<li>Check OD600 to make sure culture is in the log phase (0.5-1) after 4-5 h of growth. Depending on the growth of the strain, make adjustments to the growth time to allow for at least two doublings. Add to 2% galactose and grow overnight in 30 &#176;C shaker. 
	NOTE: The OD600 after the overnight growth may vary but culture should be saturated. Note that cells do not need to be removed from 0.1% glucose before addition of 2% galactose for this growth protocol. We recommend harvesting 3 OD of uninduced and induced cells at this step for SDS-PAGE and western blot for verification (Figure 3A,B, Lane 1-2).</li>
	<li>Harvest cells by centrifugation at 4500 x g for 15 min. A swinging bucket rotor that accommodates the 1 L volume culture may be used here.</li>
	<li>Transfer the yeast pellet into a 50 mL conical tube; a second centrifugation step may be performed as needed. The cell pellet will generally be around 10-15 mL in volume as measured by graduation markings and may be used immediately or stored at -80 &#176;C.
	<ol>
		<li>Resuspend pellet in 15 mL purification buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1.5 mM MgCl2, 1 mM Dithiothreitol (DTT), protease inhibitor cocktail) to make final volume around 30 mL.</li>
	</ol>
	</li>
	<li>Chill a homogenizer 4 &#176;C before use and equilibrate it with purification buffer. Lyse cells using a mechanical cell disruptor or homogenizer. Load sample into the homogenizer and lyse at 20,000-25,000 psi for 2-3 rounds; note that as cells become lysed, more input pressure is required to maintain 20,000-25,000 psi. Collect the cell lysate in a 50 mL conical tube on ice. 
	NOTE: If additional samples need to be lysed, the homogenizer should be thoroughly cleaned and equilibrated before loading the next sample. Keep all cell lysates on ice.</li>
	<li>Immediately clear the cell lysate at 35,000 x g for 1 h at 4 &#176;C. Carefully transfer the supernatant into new tube. Note that the lipids from cell lysis will float to the top during centrifugation and will not affect purification. 
	NOTE: We recommend saving 0.5-1% of the lysate and pellet for SDS-PAGE samples. Typically, no major differences are observed and an additional western blot may be done to confirm TAP protein solubility (Figure 4, Lanes 1 and 2, respectively).</li>
	<li>Equilibrate 300 &#181;L of IgG sepharose beads with purification buffer. Add to the cleared cell lysate and incubate for 2 h, rotating at 4 &#176;C.</li>
	<li>Collect beads in a 10 mL chromatography column and allow unbound lysate to flow through.</li>
	<li>Wash beads using 10 mL of wash buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1.5 mM MgCl2, 1 mM DTT), adding 1 mL at a time and allowing it to flow through completely. 
	NOTE: We recommend saving 2% of the bound-IgG beads for SDS-PAGE sample. Typically, we observe four major bands; two SNX-BAR proteins and IgG Heavy and Light Chains (Figure 4, Lane 3). We recommend saving equivalent amounts of &#39;eluate&#39; and &#39;IgG beads after TEV&#39; to compare for TEV cleavage efficiency.</li>
	<li>Collect beads and transfer them to a microcentrifuge tube. Add to 500 &#181;L of total volume with fresh wash buffer and 2 &#181;L of 10 mg/mL TEV protease and incubate overnight, rotating at 4 &#176;C.</li>
	<li>Next morning, remove the supernatant completely using a 27 G needle and assess protein purity by 10% polyacrylamide SDS-PAGE (Figure 4, Lane 4). 
	NOTE: We typically obtain 500 &#181;L of 0.5-1 mg/mL of 95% pure heterodimer (Figure 4, Lane 4). Additional purification using calmodulin resin can also be done, however we typically see a significant reduction in yield and recommend stopping here if purity is &#62;90%. TEV does not interfere with liposome binding assays, though TEV can be additionally removed using Ni-NTA agarose beads15.</li>
	<li>To concentrate, transfer the sample to a 0.5 mL centrifugal filter with 10 KDa cutoff and centrifuge according to manufacturer&#39;s instructions to 50 &#181;L or less. Quantify the concentrated proteins using the Bradford protein assay. Store at 4 &#176;C and use within one week.</li>
</ol>
3. Liposome Preparation
<ol>
	<li>Purchase commercially available lipids: phosphatidylserine (PS), PI3P, ergosterol, and phosphatidylcholine (PC). If needed, resuspend in recommended solvent to make stock lipids. 
	NOTE: Lipids are resuspended in a methanol/chloroform mixture per manufacturer&#39;s recommendations. Make sure lipid stocks are clear and warmed to room temperature before using. Resuspended lipids can be stored under argon gas and sealed using wax film (or equivalent) at -20 &#176;C for 6-12 months or until loss of activity is observed.</li>
	<li>Calculate the volume required of each lipid stock to create a mixture with the desired lipid composition (see Table 1). Assume a total of 1 mole of lipids in the lipid mixture.</li>
	<li>Perform this step in a chemical fume hood. Clean glass syringes by drawing up a full syringe volume of chloroform and discarding it in a waste container. Repeat two more times. When transferring chloroform, use only glass syringes or pipettes. When drawing up chloroform, pull on the stopper slowly to prevent introduction of gas bubbles into the syringe.</li>
	<li>Use glass syringes to transfer stock lipids as calculated into a clean glass culture tube to make a final lipid mixture of 1% PI3P, 20% ergosterol, 30% PS, PC (Table 1). Depending on the solvents each lipid is resuspended in, the mixture may turn cloudy upon addition of each lipid. 
	NOTE: To vary concentrations of PS (0-30%), adjust volumes accordingly and compensate with varying PC (Table 1).</li>
	<li>Carefully dry down the lipid mixture using nitrogen gas directed at the lipid mixture in a circular motion to dry lipids uniformly. Use low gas flow to keep the lipids at the bottom of the glass tube during the drying process. Wrap the glass culture tube with foil, leaving the opening uncovered, and further dehydrate in vacuum for 1 h.</li>
	<li>Add 400 &#181;L of binding buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1 mM MgCl2) to completely dehydrate lipids to make a final liposome concentration of 2.5 mM. 
	NOTE: Final liposome concentration can be adjusted by adding more or less binding buffer. For instance, 200 &#181;L binding buffer may be added to produce a liposome concentration of 5 mM if needed (see below).
	<ol>
		<li>Resuspend lipids by shaking on medium speed on a vortex at room temperature for 30 min. Buffer should appear cloudy as lipids are resuspended.</li>
	</ol>
	</li>
	<li>Transfer resuspended liposomes to a microcentrifuge tube. From this point on, plastic pipette tips may be used as lipids are no longer resuspended in chloroform. Note that the liposome solution should look cloudy.</li>
	<li>Freeze-thaw liposomes seven to eight times by submerging the microcentrifuge tube first in liquid nitrogen, then in a 37 &#176;C water bath. The liposome mixture should appear completely frozen and solid by eye before thawing.</li>
	<li>Perform steps involving chloroform in a chemical fume hood. Clean two 1 mL glass syringes by drawing up and discarding full syringe volumes of chloroform, three times each, to remove any residual lipids. Equilibrate each glass syringe with ultrapure water by drawing up two syringe volumes, then equilibrate with binding buffer by drawing up two syringe volumes.</li>
	<li>Assemble the mini-extruder according to manufacturer&#39;s recommendations. Equilibrate one 200 nm membrane and two pieces of filter supports (see Table of Materials) by submerging each in binding buffer.</li>
	<li>Sandwich the membrane between the filter supports and place in the mini-extruder. To reduce the dead volume in the assembled mini-extruder and to make sure the assembly is air-tight, pass a volume of binding buffer comparable to the volume of the liposome mixture through mini-extruder using the 1 mL glass syringes.</li>
	<li>Use one of the 1 mL glass syringes and draw up the liposome mixture. Invert the microcentrifuge tube to collect the last of the liposome mixture in the tube cap for drawing up into the glass syringe.</li>
	<li>Extrude liposomes by passing through the 200 nm membrane 19-21 times. Collect extruded liposomes in new a microcentrifuge tube. 
	NOTE: Extruded liposomes should appear less cloudy than liposomes before extrusion. Liposomes should be used the same day and stored on ice. The last extrusion should place liposomes in the syringe opposite of the one it began in.</li>
</ol>
4. SNX-BAR Liposome Binding and Tubulation
<ol>
	<li>Preclear purified protein at 100,000 x g in an ultracentrifuge for 20 min at 4 &#176;C prior to conducting liposome binding and sedimentation experiments. Remove the supernatant and transfer it to a new microcentrifuge tube; do not disturb the pellet if there is one.</li>
	<li>To perform liposome binding and tubulation assays, incubate 4 &#181;M purified Snx4-Atg20 and 2.5 mM liposomes in a total reaction volume of 20 &#181;L, varying the volume of liposomes added. 
	NOTE: In the same experiment, the same volume of liposomes should be used.</li>
	<li>Incubate the reaction at 30 &#176;C for 30 min. 
	NOTE: We suggest maximizing the amount of 2.5 mM liposomes added to each reaction by using no less than 10 &#181;L liposomes to allow for visualization during sedimentation. If 10 &#181;L of 2.5 mM liposomes is used in a 20 &#181;L reaction, the final concentration of liposomes will be 1.25 mM. If the purified proteins are dilute and more volume is required for 4 &#181;M protein, the lipids may be resuspended in 200 &#181;L during the rehydration step to double the concentration of liposomes (see Step 3.6); however, this will require knowing the protein concentration prior to making the liposomes.</li>
	<li>Visualize and quantify liposome tubulation.
	<ol>
		<li>Process liposome binding reactions immediately for electron microscopy analysis. Spot samples onto a carbon-coated copper mesh grid and negative stain using 1% uranyl acetate (Figure 5B)16.</li>
		<li>Analyze samples on a transmission electron microscope (200 kV).</li>
		<li>Use image analysis software to measure and quantify tubule diameter. To accurately quantify tubule diameter of a single tubule, take three diameter measurements along the length of a tubule and average (Figure 5B). 
		NOTE: Two-way analysis of variance was used to determine statistical significance (Figure 5E). Uranyl acetate is both radioactive and toxic. Proper lab safety certification is required to perform this step.</li>
	</ol>
	</li>
	<li>Liposome binding and sedimentation.
	<ol>
		<li>Transfer the reaction (20 &#181;L, from step 4.3) to a polycarbonate centrifuge tube and use a compatible rotor to spin at 100,000 x g in an ultracentrifuge for 20 min at 4 &#176;C. Carefully remove the supernatant and transfer to new microcentrifuge tube. Note that the pellet should stay intact.</li>
		<li>Resuspend the pellet in SDS-PAGE 40 &#181;L of sample buffer and transfer to a new microcentrifuge tube. Add 20 &#181;L of sample buffer to the supernatant. Load equivalent amounts of pellet and supernatant in a 10% polyacrylamide SDS-PAGE gel and perform Coomassie staining to visualize SNX-BARs bound to liposomes (Figure 5A).</li>
		<li>To quantify the amount of SNX-BAR complex in the pellet fraction, quantify band intensities using densitometry and quantify the proportion of SNX-BAR proteins in the pellet fraction. 
		NOTE: Two-way analysis of variance was used to determine statistical significance (Figure 5C).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here, we demonstrate an optimized workflow to purify SNX-BAR dimers in yeast and two assays to evaluate their biophysical properties on synthetic liposomes. The main advantage over typical recombinant protein expression in Escherichia coli or other systems is the ability to evenly express SNX-BAR proteins in a native host, thus avoiding the toxicity and insolubility issues found in purifying SNX-BARs in other systems. It is also notable that our system does not require molecular cloning or the harboring of multi...

Membrane-bound organelles such as the plasma membrane, the endoplasmic reticulum, the Golgi apparatus, lysosome (yeast vacuole), and endosome comprise the endomembrane system of the eukaryotic cell. Most organelles have the ability to communicate and exchange material with other organelles through vesicle transport carriers. How the cell coordinates the packaging and formation of vesicle transport carriers within the endomembrane system is not well understood. However, the proteins and lipids that constitute much of the endomembrane system are known to originate from internalizing endocytic vesicles from the plasma membrane (PM). The endosome is the primary acceptor organelle for these vesicles and is comprised of multiple interconnected sets of tubular organelles. The principal function of the endosome is to facilitate nutrient acquisition, regulate protein and lipid turnover, protect from pathogen infection, and to serve as the primary replenishing source of lipids for the plasma membrane. As the endosome receives the bulk of cargo proteins and lipids from the plasma membrane, it acts as a sorting compartment by isolating cargos into tubular endosomal transport carriers (ETCs). Any proteins not sequestered into ETCs are left to be degraded via the endo-lysosomal system. The dysregulation of cargo sorting into ETCs can lead to the loss of nutrient uptake, protein turnover or lipid homeostasis, resulting in numerous metabolic, developmental, and neurological disorders1,2. However, despite ETCs central role at the endosome, the underlying mechanism of how the endosome can selectively coordinate the packaging of a multitude of heterogeneous cargos into tubular carriers is not known.
The sorting nexin (SNX) family is an evolutionarily conserved class of proteins that have been found to be critical for many vesicle transport reactions in the cell3,4,5. Sorting nexins are recruited to the endosome membrane and aid in cargo capture via their characteristic phox homology (PX) domain, which binds phosphatidylinositol-3-monophosphate (PtdIns(3)P), a lipid enriched on the endosome membrane. Mammals encode thirty-three SNX proteins, which can be further divided into multiple subfamilies, according to the presence of other domains1. Most notably, the SNX-BAR subfamily is the largest subfamily consisting of twelve in human, while in budding yeast, Saccharomyces cerevisiae, the subfamily is reduced to just seven SNX-BARs. SNX-BAR proteins have both a PX domain and a Bin-Amphiphysin-Rvs (BAR) domain that triggers lipid reservoirs to bind positive curvature membranes. Consequently, the SNX-BAR family has a natural affinity for the endosome and can mediate ETC formation via their membrane remodeling abilities. In vitro, the remodeling properties of SNX-BARs can be induced by the addition of purified SNX-BARs to synthetic liposomes and the subsequent formation of narrow, coated tubules can be visualized by electron microscopy. Using these methods, researchers have determined that both oligomerization concentration and constriction strength appear to vary amongst the SNX-BAR family suggesting they could aid in both the formation and scission of ETCs.
The SNX-BARs can be further classified by their exclusive dimerization properties. In vitro binding assays and structural studies have demonstrated that SNX-BAR proteins can only form specific homodimers or heterodimers. Therefore, in principle, each potential SNX-BAR dimer-oligomer could provide a tubule coat for a cargo-specific trafficking pathway and likewise, the restricted oligomerization of the other SNX-BAR protomers, can also define distinct export pathways. However, due to the large number of SNX-BARs and diversity within the SNX family, a one sorting nexin-one cargo hypothesis is highly unlikely. Instead a coordinated effort using a multitude of factors such as SNX-BARs, cargo, lipid specificity and other dependencies is more probable. Likewise, recent studies of the yeast SNX4 family revealed evidence for additional lipid specificity, beyond PtdIns(3)P, to potentiate endosome transport carriers6. In this study, SNX-BAR homodimer Mvp1-Mvp1 was purified from bacteria and native heterodimers Snx4-Atg20 and Vps5-Vps17 were expressed and purified in high yield from yeast, while only Snx4-Atg20 was found to preferentially bind phosphatidylserine (PS) and form narrow tube-like structures in liposome binding studies6. While others in the field have revealed important properties of the SNX-BARs using recombinantly purified SNX-BAR homodimers from bacteria, toxicity associated with expressing SNX-BAR heterodimers in similar systems have hindered their native characterization7,8,9,10. Therefore, without a reliable system to obtain pure recombinantly expressed native heterodimers, researchers must forgo these lines of investigation. In Figure 1, we present a four-part workflow to 1) construct a yeast strain overexpressing SNX-BAR heterodimers for tandem affinity purification, 2) express and purify native SNX-BAR heterodimers, 3) prepare unilamellar synthetic liposomes, and 4) set up a liposome tubulation or sedimentations assay, providing a vital tool for researchers to investigate the growing catalogue of sorting nexins found in nature.

<table><tbody><tr><td>0.2 &amp;mu;m PC Membranes</td><td>Avanti</td><td>610006</td><td></td></tr><tr><td>10 mL Poly-Prep Chromatography column (Bio-Rad)</td><td>Bio-Rad</td><td>731-1550</td><td></td></tr><tr><td>27 G needle</td><td>BD Biosciences</td><td>301629</td><td></td></tr><tr><td>Amicon Ultra Centrifugal Filter with 10K cutoff</td><td>Amicon</td><td>UFC501024</td><td></td></tr><tr><td>Avestin EmulsiFlex-C3 Homogenizer</td><td>Avestin</td><td>EF-C3</td><td></td></tr><tr><td>BCA assay</td><td>Pierce</td><td>23225</td><td></td></tr><tr><td>Beckman Optima MAX-XP Ultracentrifuge</td><td>Beckman Coulter</td><td>393315</td><td></td></tr><tr><td>Complete Protease Inhibitor Cocktail</td><td>Roche</td><td>4693116001</td><td></td></tr><tr><td>DOPC</td><td>Avanti</td><td>850375</td><td></td></tr><tr><td>DOPS</td><td>Avanti</td><td>840035</td><td></td></tr><tr><td>Ergosterol (Sigma)</td><td>Sigma</td><td>47130-U</td><td></td></tr><tr><td>Extruder Set with Block 0.2 &amp;mu;L/1 mL</td><td>Avanti</td><td>610000</td><td></td></tr><tr><td>FEI Tecnai F20 transmission electron microscope (200 kV)</td><td></td><td></td><td></td></tr><tr><td>Glass culture tubes</td><td>VWR</td><td>47729-570</td><td></td></tr><tr><td>IgG sepharose beads (GE Healthcare)</td><td>GE Healthcare</td><td>17-0969-01</td><td></td></tr><tr><td>Microlter glass syringes</td><td>Hamilton</td><td>7637-01</td><td></td></tr><tr><td>New Brunswick Excella E25</td><td>Eppendorf</td><td>M1353-0000</td><td>or equivalent shaking 30 &amp;deg;C</td></tr><tr><td>Ni-NTA Magnetic Agarose Beads</td><td>Pierce</td><td>78605</td><td></td></tr><tr><td>Optima XE-90 Ultracentrifuge</td><td>Beckman Coulter</td><td>A94516</td><td></td></tr><tr><td>Parafilm M</td><td>VWR</td><td>52858-076</td><td></td></tr><tr><td>PI3P</td><td>Echelon</td><td>P-3016</td><td>or Echelon equivalent</td></tr><tr><td>Polycarbonate bottle assembly</td><td>Beckman Coulter</td><td>355622</td><td></td></tr><tr><td>TLA-100 Fixed-Angle Rotor</td><td>Beckman Coulter</td><td>343840</td><td></td></tr><tr><td>Type 45 Ti Rotor</td><td>Beckman Coulter</td><td></td><td></td></tr><tr><td>Vacuum Desiccator, Bottom and Lid with Socket Valve</td><td>VWR</td><td>75871-436</td><td></td></tr><tr><td>Vacuum Pump Alcatel (Pascal 2005 C1)</td><td>A&amp;amp;J Vacuum</td><td>PN07050</td><td></td></tr><tr><td>Vortex with foam holder</td><td>VWR</td><td>10153-838</td><td></td></tr><tr><td>VWR KIT MICROTUBE</td><td>VWR</td><td>12620-880</td><td></td></tr></tbody></table>

expression, purification, and liposome binding of budding yeast snx-bar heterodimers

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and lipids during endocytosis, sorting within the endosomal system, and autophagy. Central to SNX-BAR protein function is the ability to form homodimers or heterodimers that bind membranes using highly conserved phox-homology (PX) and BAR (Bin/Amphiphysin/Rvs) domains. In addition, oligomerization of SNX-BAR dimers on membranes can elicit the formation of membrane tubules and vesicles and this activity is thought to reflect their functions as coat proteins for endosome-derived transport carriers. Researchers have long utilized in vitro binding studies using recombinant SNX-BAR proteins on synthetic liposomes or giant unilamellar vesicles (GUVs) to reveal the precise makeup of lipids needed to drive membrane remodeling, thus revealing their mechanism of action. However, due to technical challenges with dual expression systems, toxicity of SNX-BAR protein expression in bacteria, and poor solubility of individual SNX-BAR proteins, most studies to date have examined SNX-BAR homodimers, including non-physiological dimers that form during expression in bacteria. Recently, we have optimized a protocol to overcome the major shortcomings of a typical bacterial expression system. Using this workflow, we demonstrate how to successfully express and purify large amounts of SNX-BAR heterodimers and how to reconstitute them on synthetic liposomes for binding and tubulation assays.

This protocol describes a method for reproducible and robust production of endogenous yeast SNX-BAR complexes that can be used for downstream membrane remodeling assays (Figure 1). The construction of the yeast strain used for purification takes advantage of the efficiency of homologous recombination in budding yeast, allowing for modifications at the genomic loci of the targeted SNX-BARs (Figure 2). This design has two advantage...

Watch this Scientific Journal Video about Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers at JoVE.com

Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

Here, we present a workflow for the expression, purification and liposome binding of SNX-BAR heterodimers in yeast.

SNX-BAR proteins are an evolutionarily conserved class of membrane remodeling proteins that play key roles in sorting and trafficking of protein and ...

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Research

JoVE Journal

Genetics

7.7K Views.  Yale School of Medicine. Here, we present a workflow for the expression, purification and liposome binding of SNX-BAR heterodimers in yeast.

Video: Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

Single Molecule Fluorescence In Situ Hybridization (smFISH) Analysis in Budding Yeast Vegetative Growth and Meiosis

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect and count individual RNA molecules. Complementary to deep sequencing-based methods, smFISH provides information about the cell-to-cell variation in transcript abundance and the subcellular localization of a given RNA. Recently, we have used smFISH to study the expression of the gene&#160;NDC80&#160;during meiosis in budding yeast, in which two transcript isoforms exist and the short transcript isoform has its entire sequence shared with the long isoform. To confidently identify each transcript isoform, we optimized known smFISH protocols and obtained high consistency and quality of smFISH data for the samples acquired during budding yeast meiosis. Here, we describe this optimized protocol, the criteria that we use to determine whether high quality of smFISH data is obtained, and some tips for implementing this protocol in&#160;other yeast strains and growth conditions.

Single Molecule Fluorescence In Situ Hybridization smFISH Analysis in Budding Yeast Vegetative Growth and Meiosis

This single molecule fluorescence in situ hybridization protocol is optimized to quantify the number of RNA molecules in budding yeast during vegetative growth and meiosis.

Single molecule fluorescence in situ hybridization (smFISH) is a powerful technique to study gene expression in single cells due to its ability to detect ...

Expression, Solubilization, and Purification of Eukaryotic Borate Transporters

The Solute Carrier 4 (SLC4) family of proteins is called the bicarbonate transporters and includes the archetypal protein Anion Exchanger 1 (AE1, also known as Band 3), the most abundant membrane protein in the red blood cells. The SLC4 family is homologous with borate transporters, which have been characterized in plants and fungi. It remains a significant technical challenge to express and purify membrane transport proteins to homogeneity in quantities suitable for&#160;structural or functional studies. Here we describe detailed procedures for the overexpression of borate transporters in Saccharomyces cerevisiae, isolation of yeast membranes, solubilization of protein by detergent, and purification of borate transporter homologs from S. cerevisiae, Arabidopsis thaliana, and Oryza sativa. We also detail a glutaraldehyde cross-linking experiment to assay multimerization of homomeric transporters. Our generalized procedures can be applied to all three proteins and have been optimized for efficacy. Many of the strategies developed here can be utilized for the study of other challenging membrane proteins.

Here we present a protocol to express, solubilize, and purify several eukaryotic borate transporters with homology to the SLC4 transporter family using yeast. We also describe a chemical cross-linking assay to assess the purified homomeric proteins for multimeric assembly. These protocols can be adapted for other challenging membrane proteins.

The Solute Carrier 4 (SLC4) family of proteins is called the bicarbonate transporters and includes the archetypal protein Anion Exchanger 1 (AE1, also ...

Biochemistry

ACT1-CUP1 Assays Determine the Substrate-Specific Sensitivities of Spliceosomal Mutants in Budding Yeast

Mutations introduced in the spliceosome or its substrate have significantly contributed to our understanding of the intricacies of spliceosomal function. Whether disease-related or functionally selected, many of these mutations have been studied using growth assays in the model organism Saccharomyces cerevisiae (yeast). The splicing-specific copper growth assay, or ACT1-CUP1 assay, provides a comprehensive analysis of mutation at the phenotypic level. The ACT1-CUP1 assay utilizes reporters that confer copper tolerance when correctly spliced. Thus, in the presence of copper, changes in yeast viability correlate to changes in mRNA production through splicing. In a typical experiment, the yeast spliceosome is challenged with different non-consensus splicing reporters and the splicing factor mutation of interest to detect any synergetic or antithetical impact on splicing. Here a full description of copper plate preparation, plating of yeast cells, and data evaluation are given. A selection of complimentary experiments is described, highlighting the versatility of the ACT1-CUP1 reporters. The ACT1-CUP1 assay is a handy tool in the splicing toolbox thanks to the direct read-out of mutational effect(s) and the comparative possibilities from the continuing use in the field.

The ACT1-CUP1 assay, a copper growth assay, provides a quick readout of precursor messenger RNA (pre-mRNA) splicing and the impact mutant splicing factors have on spliceosomal function. This study provides a protocol and highlights the customization possible to address the splicing question of interest.

Mutations introduced in the spliceosome or its substrate have significantly contributed to our understanding of the intricacies of spliceosomal function. ...

High-Speed Magnetic Tweezers for Nanomechanical Measurements on Force-Sensitive Elements

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are therefore poised to be useful in the field of mechanobiology. Since the method commonly relies on image-based tracking of magnetic beads, the speed limit in recording and analyzing images, as well as the thermal fluctuations of the beads, has long hampered its application in observing small and fast structural changes in target molecules. This article describes detailed methods for the construction and operation of a high-resolution MT setup that can resolve nanoscale, millisecond dynamics of biomolecules and their complexes. As application examples, experiments with DNA hairpins and SNARE complexes (membrane-fusion machinery) are demonstrated, focusing on how their transient states and transitions can be detected in the presence of piconewton-scale forces. We expect that high-speed MTs will continue to enable high-precision nanomechanical measurements on molecules that sense, transmit, and generate forces in cells, and thereby deepen our molecular-level understanding of mechanobiology.

Here, we describe a high-speed magnetic tweezer setup that performs nanomechanical measurements on force-sensitive biomolecules at the maximum rate of 1.2 kHz. We introduce its application to DNA hairpins and SNARE complexes as model systems, but it will be also applicable to other molecules involved in mechanobiological events.

Single-molecule magnetic tweezers (MTs) have served as powerful tools to forcefully interrogate biomolecules, such as nucleic acids and proteins, and are ...

Locus-Specific Chromatin Isolation from Yeast Cells

Single-Copy Gene Locus Chromatin Purification in Saccharomyces cerevisiae

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone octamer. Chromatin is defined as the entity of NCPs and numerous other protein complexes, including transcription factors, chromatin remodeling and modifying enzymes. It is still unclear how these protein-DNA interactions are orchestrated at the level of specific genomic loci during different stages of the cell cycle. This is mainly due to the current technical limitations, which make it challenging to obtain precise measurements of such dynamic interactions. Here, we describe an improved method combining site-specific recombination with an efficient single-step affinity purification protocol to isolate a single-copy gene locus of interest in its native chromatin state. The method allows for the robust enrichment of the target locus over genomic chromatin, making this technique an effective strategy for identifying and quantifying protein interactions in an unbiased and systematic manner, for example by mass spectrometry. Further to such compositional analyses, native chromatin purified by this method likely reflects the in vivo situation regarding nucleosome positioning and histone modifications and is, therefore, amenable to further structural and biochemical analyses of chromatin derived from virtually any genomic locus in yeast.

Author Spotlight: Advancing Chromatin Research and Overcoming Limitations with a High-Enrichment Locus-Specific Chromatin Isolation Protocol 

This protocol presents a locus-specific chromatin isolation method based on site-specific recombination to purify a single-copy gene locus of interest in its native chromatin context from budding yeast, Saccharomyces cerevisiae.

The basic organizational unit of eukaryotic chromatin is the nucleosome core particle (NCP), which comprises DNA wrapped ~1.7 times around a histone ...

Budding Yeast Protein Extraction and Purification for the Study of Function, Interactions, and Post-translational Modifications

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously hard to disrupt. Here we describe the use of a bead mill homogenizer for the extraction of proteins into buffers optimized to maintain the functions, interactions and post-translational modifications of proteins. Logarithmically growing cells expressing the protein of interest are grown in a liquid growth media of choice. The growth media may be supplemented with reagents to induce protein expression from inducible promoters (e.g. galactose), synchronize cell cycle stage (e.g. nocodazole), or inhibit proteasome function (e.g. MG132). Cells are then pelleted and resuspended in a suitable buffer containing protease and/or phosphatase inhibitors and are either processed immediately or frozen in liquid nitrogen for later use. Homogenization is accomplished by six cycles of 20 sec&#160;bead-beating (5.5 m/sec), each followed by one minute incubation on ice. The resulting homogenate is cleared by centrifugation and small particulates can be removed by filtration. The resulting cleared whole cell extract (WCE) is precipitated using 20% TCA for direct analysis of total proteins by SDS-PAGE followed by Western blotting. Extracts are also suitable for affinity purification of specific proteins, the detection of post-translational modifications, or the analysis of co-purifying proteins. As is the case for most protein purification protocols, some enzymes and proteins may require unique conditions or buffer compositions for their purification and others may be unstable or insoluble under the conditions stated. In the latter case, the protocol presented may provide a useful starting point to empirically determine the best bead-beating strategy for protein extraction and purification. We show the extraction and purification of an epitope-tagged SUMO E3 ligase, Siz1, a cell cycle regulated protein that becomes both sumoylated and phosphorylated, as well as a SUMO-targeted ubiquitin ligase subunit, Slx5.

The preparation of high quality yeast cell extracts is a necessary first step in the analysis of individual proteins or entire proteomes. Here we describe a fast, efficient, and reliable homogenization protocol for budding yeast cells that has been optimized to preserve protein functions, interactions, and post-translational modifications.

Homogenization by bead beating is a fast and efficient way to release DNA, RNA, proteins, and metabolites from budding yeast cells, which are notoriously ...

Biology

Protein Purification Technique that Allows Detection of Sumoylation and Ubiquitination of Budding Yeast Kinetochore Proteins Ndc10 and Ndc80

Post-translational Modifications (PTMs), such as phosphorylation, methylation, acetylation, ubiquitination, and sumoylation, regulate the cellular function of many proteins. PTMs of kinetochore proteins that associate with centromeric DNA mediate faithful chromosome segregation to maintain genome stability. Biochemical approaches such as mass spectrometry and western blot analysis are most commonly used for identification of PTMs. Here, a protein purification method is described that allows the detection of both sumoylation and ubiquitination of the kinetochore proteins, Ndc10 and Ndc80, in Saccharomyces cerevisiae. A strain that expresses polyhistidine-Flag-tagged Smt3 (HF-Smt3) and Myc-tagged Ndc10 or Ndc80 was constructed and used for our studies. For detection of sumoylation, we devised a protocol to affinity purify His-tagged sumoylated proteins by using nickel beads and used western blot analysis with anti-Myc antibody to detect sumoylated Ndc10 and Ndc80. For detection of ubiquitination, we devised a protocol for immunoprecipitation of Myc-tagged proteins and used western blot analysis with anti-Ub antibody to show that Ndc10 and Ndc80 are ubiquitinated. Our results show that epitope tagged-protein of interest in the His-Flag tagged Smt3 strain facilitates the detection of multiple PTMs. Future studies should allow exploitation of this technique to identify and characterize protein interactions that are dependent on a specific PTM.

This manuscript describes the detection of sumoylation and ubiquitination of kinetochore proteins, Ndc10 and Ndc80, in the budding yeast Saccharomyces cerevisiae.

Post-translational Modifications (PTMs), such as phosphorylation, methylation, acetylation, ubiquitination, and sumoylation, regulate the cellular ...

Microscopy of Fission Yeast Sexual Lifecycle

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the mechanics of cell division and cell polarity. Furthermore, classical experiments on its sexual reproduction have yielded results pivotal to current understanding of DNA recombination and meiosis. More recent analysis of fission yeast mating has raised interesting questions on extrinsic stimuli response mechanisms, polarized cell growth and cell-cell fusion. To study these topics in detail we have developed a simple protocol for microscopy of the entire sexual lifecycle. The method described here is easily adjusted to study specific mating stages. Briefly, after being grown to exponential phase in a nitrogen-rich medium, cell cultures are shifted to a nitrogen-deprived medium for periods of time suited to the stage of the sexual lifecycle that will be explored. Cells are then mounted on custom, easily built agarose pad chambers for imaging. This approach allows cells to be monitored from the onset of mating to the final formation of spores.

We provide a reproducible basic method for the long-term microscopy of the fission yeast sexual lifecycle. With minor adjustments described, the presented protocol allows research focus on different steps of the reproductive process.

The fission yeast Schizosaccharomyces pombe has been an invaluable model system in studying the regulation of the mitotic cell cycle progression, the ...

Purification of Native Complexes for Structural Study Using a Tandem Affinity Tag Method

Affinity purification approaches have been successful in isolating native complexes for proteomic characterization. Structural heterogeneity and a degree of compositional heterogeneity of a complex do not usually impede progress in conducting such studies. In contrast, a complex intended for structural characterization should be purified in a state that is both compositionally and structurally homogeneous as well as at a higher concentration than required for proteomics. Recently, there have been significant advances in the application of electron microscopy for structure determination of large macromolecular complexes. This has heightened interest in approaches to purify native complexes of sufficient quality and quantity for structural determination by electron microscopy. The Tandem Affinity Purification (TAP) method has been optimized to extract and purify an 18-subunit, ~ 0.8 MDa ribonucleoprotein assembly from budding yeast (Saccharomyces cerevisiae) suitable for negative stain and electron cryo microscopy. Herein is detailed the modifications made to the TAP method, the rationale for making these changes, and the approaches taken to assay for a compositionally and structurally homogeneous complex.

The Tandem Affinity Purification (TAP) method has been used extensively to isolate native complexes from cellular extract, primarily eukaryotic, for proteomics. Here, we present a TAP method protocol optimized for purification of native complexes for structural studies.

Affinity purification approaches have been successful in isolating native complexes for proteomic characterization. Structural heterogeneity and a degree ...

Applications of pHluorin for Quantitative, Kinetic and High-throughput Analysis of Endocytosis in Budding Yeast

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal remodeling and vesicular trafficking in living cells. Quantitative methodologies using chimeric GFP fusions have been developed for many applications; however, GFP is somewhat resistant to proteolysis, thus its fluorescence persists in the lysosome/vacuole, which can impede quantification of cargo trafficking in the endocytic pathway. An alternative method for quantifying endocytosis and post-endocytic trafficking events makes use of superecliptic pHluorin, a pH-sensitive variant of GFP that is quenched in acidic environments. Chimeric fusion of pHluorin to the cytoplasmic tail of transmembrane cargo proteins results in a dampening of fluorescence upon incorporation of the cargo into multivesicular bodies (MVBs) and delivery to the lysosome/vacuole lumen. Thus, quenching of vacuolar fluorescence facilitates quantification of endocytosis and early events in the endocytic pathway. This paper describes methods using pHluorin-tagged cargos for quantification of endocytosis via fluorescence microscopy, as well as population-based assays using flow cytometry.

Accurate quantification of vesicular trafficking events often provides key insights into roles for specific proteins and the effects of mutations. This paper presents methods for using superecliptic pHluorin, a pH-sensitive GFP variant, as a tool for quantification of endocytic events in living cells using quantitative fluorescence microscopy and flow cytometry.

Green fluorescent protein (GFP) and its variants are widely used tools for studying protein localization and dynamics of events such as cytoskeletal ...