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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 micrometer 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 Gauge 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 microlter/1mL</td>
			<td>Avanti</td>
			<td>610000</td>
			<td></td>
		</tr>
		<tr>
			<td colspan="2">FEI Tecnai F20 transmission electron microscope (200 kV)</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 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&#38;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 ...

expression-purification-liposome-binding-budding-yeast-snx-bar

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

Genetic Engineering of an Unconventional Yeast for Renewable Biofuel and Biochemical Production

Yarrowia lipolytica is a non-pathogenic, dimorphic and strictly aerobic yeast species. Owing to its distinctive physiological features and metabolic characteristics, this unconventional yeast is not only a good model for the study of the fundamental nature of fungal differentiation but is also a promising microbial platform for biochemical production and various biotechnological applications, which require extensive genetic manipulations. However, genetic manipulations of Y. lipolytica have been limited due to the lack of an efficient and stable genetic transformation system as well as very high rates of non-homologous recombination that can be mainly attributed to the KU70 gene. Here, we report an easy and rapid protocol for the efficient genetic transformation and for gene deletion in Y. lipolytica Po1g. First, a protocol for the efficient transformation of exogenous DNA into Y. lipolytica Po1g was established. Second, to achieve the enhanced double-crossover homologous recombination rate for further deletion of target genes, the KU70 gene was deleted by transforming a disruption cassette carrying 1 kb homology arms. Third, to demonstrate the enhanced gene deletion efficiency after deletion of the KU70 gene, we individually deleted 11 target genes encoding alcohol dehydrogenase and alcohol oxidase using the same procedures on the KU70 knockout platform strain. It was observed that the rate of precise homologous recombination increased substantially from less than 0.5% for deletion of the KU70 gene in Po1g to 33%-71% for the single gene deletion of the 11 target genes in Po1g KU70&#916;. A replicative plasmid carrying the hygromycin B resistance marker and the Cre/LoxP system was constructed, and the selection marker gene in the yeast knockout strains was eventually removed by expression of Cre recombinase to facilitate multiple rounds of targeted genetic manipulations. The resulting single-gene deletion mutants have potential applications in biofuel and biochemical production.

We herein report methods on the molecular genetic manipulation of the Yarrowia lipolytica Po1g strain for improved gene deletion efficiency. The resulting engineered Y. lipolytica strains have potential applications in biofuel and biochemical production.

Yarrowia lipolytica is a non-pathogenic, dimorphic and strictly aerobic yeast species. Owing to its distinctive physiological features and metabolic ...

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting mutant alleles reside at their endogenous genomic sites and no exogenous DNA sequences are left in the genome following the procedure. Since in haploid yeast cells each of the four core histone proteins is encoded by two non-allelic genes with highly homologous open reading frames (ORFs), targeting mutagenesis specifically to one of two genes encoding a particular histone protein can be problematic. The strategy we describe here bypasses this problem by utilizing sequences outside, rather than within, the ORF of the target genes for the homologous recombination step. Another feature of this system is that the regions of DNA driving the homologous recombination steps can be made to be very extensive, thus increasing the likelihood of successful integration events. These features make this strategy particularly well-suited for histone gene mutagenesis, but can also be adapted for mutagenesis of other genes in the yeast genome.

Targeted in Situ Mutagenesis of Histone Genes in Budding Yeast

A strategy for generating mutations in histone genes at their endogenous location in Saccharomyces cerevisiae is presented.

We describe a PCR- and homologous recombination-based system for generating targeted mutations in histone genes in budding yeast cells. The resulting ...

Production and Purification of Baculovirus for Gene Therapy Application

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a promising vector for gene therapy application. Here, baculovirus is produced by transient transfection of the baculovirus plasmid DNA (bacmid) in an adherent culture of Sf9 cells. Baculovirus is subsequently expanded in Sf9 cells in a serum-free suspension culture until the desired volume is obtained. It is then purified from the culture supernatant using heparin affinity chromatography. Virus supernatant is loaded onto the heparin column which binds baculovirus particles in the supernatant due to the affinity of heparin for baculovirus envelop glycoprotein. The column is washed with a buffer to remove contaminants and baculovirus is eluted from the column with a high-salt buffer. The eluate is diluted to an isotonic salt concentration and baculovirus particles are further concentrated using ultracentrifugation. Using this method, baculovirus can be concentrated up to 500-fold with a 25% recovery of infectious particles. Although the protocol described here demonstrates the production and purification of the baculovirus from cultures up to 1 L, the method can be scaled-up in a closed-system suspension culture to produce a clinical-grade vector for gene therapy application.

In this protocol, baculovirus is produced by transient transfection of baculovirus plasmid into Sf9 cells and amplified in a serum-free suspension culture. The supernatant is purified by heparin affinity chromatography and further concentrated by ultracentrifugation. This protocol is useful for the production and purification of baculovirus for gene therapy application.

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a ...

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to abiotic stresses and diseases. As the plant microbiome can be highly complex, low-cost, high-throughput methods such as amplicon-based sequencing of the 16S rRNA gene are often preferred for characterizing its microbial composition and diversity. However, the selection of appropriate methodology when conducting such experiments is critical for reducing biases that can make analysis and comparisons between samples and studies difficult. This protocol describes in detail a standardized methodology for the collection and extraction of DNA from soil, rhizosphere, and root samples. Additionally, we highlight a well-established 16S rRNA amplicon sequencing pipeline that allows for the exploration of the composition of bacterial communities in these samples, and can easily be adapted for other marker genes. This pipeline has been validated for a variety of plant species, including sorghum, maize, wheat, strawberry, and agave, and can help overcome issues associated with the contamination from plant organelles.

Here, we describe a protocol to obtain amplicon sequence data of soil, rhizosphere, and root endosphere microbiomes. This information can be used to investigate the composition and diversity of plant-associated microbial communities, and is suitable for the use with a wide range of plant species.

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to ...

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 ...

Informatic Analysis of Sequence Data from Batch Yeast 2-Hybrid Screens

We have adapted the yeast 2-hybrid assay to simultaneously uncover dozens of transient and static protein interactions within a single screen utilizing high-throughput short-read DNA sequencing. The resulting sequence datasets can not only track what genes in a population that are enriched during selection for positive yeast 2-hybrid interactions, but also give detailed information about the relevant subdomains of proteins sufficient for interaction. Here, we describe a full suite of stand-alone software programs that allow non-experts to perform all the bioinformatics and statistical steps to process and analyze DNA sequence fastq files from a batch yeast 2-hybrid assay. The processing steps covered by these software include: 1) mapping and counting sequence reads corresponding to each candidate protein encoded within a yeast 2-hybrid prey library; 2) a statistical analysis program that evaluates the enrichment profiles; and 3) tools to examine the translational frame and position within the coding region of each enriched plasmid that encodes the interacting proteins of interest.

Deep sequencing of yeast populations selected for positive yeast 2-hybrid interactions potentially yields a wealth of information about interacting partner proteins. Here, we describe the operation of specific bioinformatics tools and customized updated software to analyze sequence data from such screens.

We have adapted the yeast 2-hybrid assay to simultaneously uncover dozens of transient and static protein interactions within a single screen utilizing ...

Mating-based Overexpression Library Screening in Yeast

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool commonly used in yeast studies. The expression of a number of neurodegenerative disease-associated proteins in yeast causes cytotoxicity and aggregate formation, recapitulating findings seen in patients with these disorders. Here, we describe a method for screening a yeast model of the Amyotrophic Lateral Sclerosis-associated protein FUS for modifiers of its toxicity. Instead of using transformation, this new screening platform relies on the mating of yeast to introduce an arrayed library of plasmids into the yeast model. The mating method has two clear advantages: first, it is highly efficient; second, the pre-transformed arrayed library of plasmids can be stored for long-term as a glycerol stock, and quickly applied to other screens without the labor-intensive step of transformation into the yeast model each time. We demonstrate how this method can successfully be used to screen for genes that modify the toxicity of FUS.

This article presents a mating-based method to facilitate overexpression screening in budding yeast using an arrayed plasmid library.

Budding yeast has been widely used as a model in studying proteins associated with human diseases. Genome-wide genetic screening is a powerful tool ...

Enhanced Yeast One-hybrid Screens To Identify Transcription Factor Binding To Human DNA Sequences

Identifying the sets of transcription factors (TFs) that regulate each human gene is a daunting task that requires integrating numerous experimental and computational approaches. One such method is the yeast one-hybrid (Y1H) assay, in which interactions between TFs and DNA regions are tested in the milieu of the yeast nucleus using reporter genes. Y1H assays involve two components: a &#8216;DNA-bait&#8217; (e.g., promoters, enhancers, silencers, etc.) and a &#8216;TF-prey,&#8217; which can be screened for reporter gene activation. Most published protocols for performing Y1H screens are based on transforming TF-prey libraries or arrays into DNA-bait yeast strains. Here, we describe a pipeline, called enhanced Y1H (eY1H) assays, where TF-DNA interactions are interrogated by mating DNA-bait strains with an arrayed collection of TF-prey strains using a high density array (HDA) robotic platform that allows screening in a 1,536 colony format. This allows for a dramatic increase in throughput (60 DNA-bait sequences against &#62;1,000 TFs takes two weeks per researcher) and reproducibility. We illustrate the different types of expected results by testing human promoter sequences against an array of 1,086 human TFs, as well as examples of issues that can arise during screens and how to troubleshoot them.

Here, we present an enhanced yeast one-hybrid screening protocol to identify the transcription factors (TFs) that can bind to a human DNA region of interest. This method uses a high-throughput screening pipeline that can interrogate the binding of &#62;1,000 TFs in a single experiment.

Identifying the sets of transcription factors (TFs) that regulate each human gene is a daunting task that requires integrating numerous experimental and ...

Exploring Sequence Space to Identify Binding Sites for Regulatory RNA-Binding Proteins

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational steps are used to regulate specific genes. Sequence-specific nucleic acid-binding proteins target specific sequences to control spatial or temporal gene expression. The binding sites in nucleic acids are typically characterized by mutational analysis. However, numerous proteins of interest have no known binding site for such characterization. Here we describe an approach to identify previously unknown binding sites for RNA-binding proteins. It involves iterative selection and amplification of sequences starting with a randomized sequence pool. Following several rounds of these steps-transcription, binding, and amplification-the enriched sequences are sequenced to identify a preferred binding site(s). Success of this approach is monitored using in vitro binding assays. Subsequently, in vitro and in vivo functional assays can be used to assess the biological relevance of the selected sequences. This approach allows identification and characterization of a previously unknown binding site(s) for any RNA-binding protein for which an assay to separate protein-bound and unbound RNAs exists.

Sequence specificity is critical for gene regulation. Regulatory proteins that recognize specific sequences are important for gene regulation. Defining functional binding sites for such proteins is a challenging biological problem. An iterative approach for identification of a binding site for an RNA-binding protein is described here and is applicable to all RNA-binding proteins.

Gene regulation plays an important role in all cells. Transcriptional, post-transcriptional (or RNA processing), translational, and post-translational ...

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. ...