Name: Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry (Video) | JoVE Journal
Uploaded: 2022-02-07T14:30:00.000+00:00
Duration: 11 min 12 s
Description: Learn organic solvent-based protein precipitation for robust proteome purification ahead of mass spectrometry. This optimized protocol maximizes recovery and purity in minutes for bottom-up and top-down workflows.

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5 minutes

This work was funded by the Natural Sciences and Engineering Research Council of Canada. The authors thank Bioinformatics Solutions Inc. (Waterloo, Canada) and SPARC BioCentre (Molecular Analysis) at the Hospital for Sick Children (Toronto, Canada) for their contributions to the acquisition of MS data.

1. Material considerations and sample pre-preparation
<ol>
	<li>Use only high purity solvents (acetone, chloroform, methanol) (&#62;99.5%) and chemicals, free of excess moisture.</li>
	<li>Prepare sodium chloride and zinc sulfate solutions (1 M) in water. 
	NOTE: Salt solutions can be stored indefinitely at room temperature, as&#160;long as they are free of contaminant or microbial growth.</li>
	<li>Use the smallest polypropylene (PP) microcentrifuge vial sufficient to retain the required volume of sample and solvents to induce precipitation.</li>
	<li>Ensure that the SDS concentration in the sample to be precipitated is no greater than 2% (w/v). If SDS is higher, dilute the sample with water.</li>
	<li>Ensure a protein concentration between 0.01 and 10 g/L for optimal precipitation efficiency. 
	NOTE: The optimal mass for precipitation ranges between 1-100 &#181;g of protein.</li>
	<li>Ensure all solvents and solutions are free of particulate matter before use. Perform either a filtration (&#60;0.5 &#181;m) or centrifugation step (10,000 x g for 1 min at room temperature) to remove undissolved particulates.</li>
	<li>If disulfide bond reduction and alkylation are required, conduct these steps prior to protein precipitation. Excess reducing and alkylating reagents will be removed through the precipitation process.</li>
	<li>Precipitate the proteins by selecting and performing one of the protocols (steps 2, 3, 4, or 5).</li>
</ol>
2. Rapid (vial-based) protein precipitation with acetone
<ol>
	<li>Pipet 90 &#181;L of (particulate-free) protein or proteome solution into a PP microcentrifuge tube. Then, add 10 &#181;L of 1 M aqueous NaCl. 
	NOTE: If the ionic strength of the proteome extract already exceeds 100 mM, no additional salt is necessary.</li>
	<li>Pipet 400 &#181;L of acetone into the sample. Cap the vial and tap the vial gently to combine the solvents. Vigorous mixing is not required. 
	NOTE: The volume of protein, salt, and acetone can be increased so long as the relative ratio of each is maintained.</li>
	<li>Allow the vial to incubate at room temperature, undisturbed, for a minimum of 2 min. 
	NOTE: Longer incubations, including those at reduced temperature (e.g., conventional acetone precipitation employs overnight precipitation in the freezer), may result in the formation of larger (visible) aggregated protein particulates (Figure 1A), which generally do not improve total protein recovery.</li>
	<li>Following incubation, place samples in a centrifuge, noting the orientation of the vial. Spin for a minimum of 2 min, at 10,000 x g or higher at room temperature.</li>
	<li>Uncap the vial and gently decant the supernatant by slowly inverting the vial to a waste container. Touch the inverted vial to a paper towel to draw residual solvent from the vial. 
	CAUTION: Waste solvents should be retained and discarded as per appropriate protocols.</li>
	<li>For SDS-containing samples, dispense 400 &#181;L of fresh acetone, being careful not to disturb the pellet. 
	NOTE: Step 2.6 is optional.
	<ol>
		<li>Immediately centrifuge the sample (10,000 x g or higher for 1 min at room temperature), placing the vial into the rotor in the same orientation as the initial spin. Decant the wash solvent as described in step 2.5.</li>
	</ol>
	</li>
	<li>Allow the sample to fully dry with the cap open (~1 min). Recap the vial and proceed with pellet solubilization (step 6).</li>
</ol>
3. Precipitation of low molecular weight (LMW) peptides (ZnSO4 + acetone)
<ol>
	<li>Dispense 54 &#181;L of proteome extract to a 2 mL PP vial, and then add 6 &#181;L of 1 M ZnSO4. 
	NOTE: Optimal recovery of LMW peptides (&#8804;5 kDa) is obtained by adding acetone to a final 97% by volume. Assuming a 2 mL PP vial, the maximal initial sample volume is 54 &#181;L.</li>
	<li>Add 1940 &#181;L of acetone to a final 97% by volume. Swirl gently to mix, and let stand undisturbed on the benchtop for a minimum of 2 min.</li>
	<li>Centrifuge (10,000 x g for 1 min at room temperature) and remove the supernatant by inverting the vial, and then touching the vial to a paper towel.</li>
	<li>For SDS-containing samples, dispense 400 &#181;L of fresh acetone, being careful not to disturb the pellet. 
	NOTE: Step 3.4 is optional.
	<ol>
		<li>Immediately centrifuge the sample as per step 3.3, placing the vial into the rotor in the same orientation as the initial spin. Decant the wash solvent as described in step 3.3.</li>
	</ol>
	</li>
	<li>Re-solubilize the resulting dry pellet in an aqueous solvent with brief vortexing or sonication (~5 min).</li>
</ol>
4. Protein precipitation by chloroform/methanol/water (CMW)
<ol>
	<li>Dispense 100 &#181;L of the protein or proteome solution into a PP vial. Add 400 &#181;L of methanol, followed by 100 &#181;L of chloroform. Cap the vial and vortex briefly to mix. 
	NOTE: For CMW precipitation, 1.5 mL vials with narrow bottoms are preferred (Figure 2A). 
	CAUTION: Chloroform solvent should be handled in an appropriate ventilation hood. All solvents that contact chloroform should be treated as halogenated waste when disposed of.</li>
	<li>Quickly dispense 300 &#181;L of water directly into the center of the vial. Cap the vial. Allow the sample to sit on the benchtop undisturbed for 1 min. 
	NOTE: The solution will immediately appear cloudy white. Avoid mixing the vial following the addition of water.</li>
	<li>Place the PP vial in a centrifuge and spin for a minimum of 5 min (10,000 x g or higher at room temperature). 
	NOTE: Once centrifuged, two visible solvent layers will form (top layer = methanol/water; bottom = chloroform). A solid protein pellet forms at the solvent interface (Figure 2A).</li>
	<li>Using a large (1 mL) micropipette tip, and holding the vial at ~45&#176;, remove ~700 &#181;L of the solvent from the upper layer at a uniform rate.</li>
	<li>Use a smaller (200 &#181;L) micropipette tip to continue removing the upper solvent layer from the ~45&#176; tilted vial. Pipet in one continuous motion until the upper solvent layer forms a bead in the vial.</li>
	<li>Add 400 &#181;L of fresh methanol to the sample vial, without disturbing the pellet, by dispensing the solvent down the side of the vial.</li>
	<li>Cap the vial. Combine the solvent layers by gently rocking the vial to swirl the solvents together. 
	NOTE: It is essential to avoid disrupting the pellet. Do not vortex the vial.</li>
	<li>Noting the orientation of the vial in the rotor, centrifuge for a minimum of 10 min (10,000 x g at room temperature). The protein pellet adheres to the bottom of the vial (Figure 2B).</li>
	<li>Tip the vial at 45&#176;, with the pellet facing down. Place the pipette tip along the upper edge of the vial and remove the supernatant with a 1 mL micropipette tip at a slow but continuous rate. Retain ~20 &#181;L of solvent in the vial.</li>
	<li>Wash the protein pellet for SDS-containing samples by slowly dispensing 400 &#181;L of fresh methanol. Do not vortex the vial.
	<ol>
		<li>Proceed directly with centrifugation (10,000 x g for 2 min at temperature), placing the vial into the rotor with the same orientation as the initial spin.</li>
	</ol>
	</li>
	<li>Remove the solvent, as per step 4.9. Allow the sample to air dry in a fumehood until the residual solvent evaporates.</li>
	<li>Consult the recommended resolubilization procedures in step 6.</li>
</ol>
5. Protein precipitation using a disposable filtration cartridge
NOTE: Each solvent-based precipitation protocol described in steps 2-5 can be performed in a two-stage filtration and extraction cartridge (see Table of Materials).
<ol>
	<li>With the plug attached to the upper filtration cartridge (Figure 3A), dispense the desired volume of the extracted proteome, salt, and solvent as outlined in one of the three options below.
	<ol>
		<li>(Option 1) For protein precipitation with acetone, combine 90 &#181;L of protein or proteome solution, 10 &#181;L of 1 M aqueous NaCl, and 400 &#181;L of acetone. Incubate for a minimum of 2 min on the benchtop. 
		NOTE: A visible precipitate will develop for concentrated protein samples (1 g/L) (Figure 3B).</li>
		<li>(Option 2) For LMW peptide precipitation, combine 15 &#181;L of the sample, 1.5 &#181;L of 1 M ZnSO4, and 485 &#181;L of acetone. Incubate for a minimum of 2 min on the benchtop. 
		NOTE: A salt concentration of 90 mM in the aqueous sample will not impact recovery relative to the 100 mM recommended in step 3.</li>
		<li>(Option 3) For CMW precipitation, add 50 &#181;L of proteome extract, 200 &#181;L of methanol, and 50 &#181;L of chloroform. Cap the vial and briefly vortex to combine.
		<ol>
			<li>Quickly dispense 150 &#181;L of water directly into the center of the vial. Incubate for 1 min on the benchtop.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Centrifuge for 2 min at 2,500 x g at room temperature with the plug still attached to the filtration cartridge.</li>
	<li>Invert the cartridge, and then unscrew and remove the plug from the cartridge base.</li>
	<li>Place the filtration cartridge in a clean vial and return to the centrifuge. Spin for 3 min at 500 x g at room temperature. Discard the flow-through solvent from the lower vial. 
	NOTE: If any solvent remains in the upper filtration cartridge, return to the centrifuge and perform an additional spin.</li>
	<li>Wash the protein pellet by adding 400 &#181;L of acetone to the filtration cartridge (for CMW precipitation, step 5.1.3, add 400 &#181;L of methanol).</li>
	<li>Centrifuge for 3 min at 500 x g at room temperature or until no solvent remains in the upper cartridge.</li>
	<li>Re-solubilize the precipitation pellet as described in step 6.</li>
</ol>
6. Resolubilization of protein pellet
<ol>
	<li>Wet the membrane at the base of the filtration cartridge by dispensing 2-5 &#181;L of isopropanol directly to the membrane immediately before the resolubilization protocols described below.</li>
	<li>Follow one of the following resolubilization methods.
	<ol>
		<li>(Option 1) Add a minimum of 20 &#181;L of aqueous buffer containing &#8805;2% SDS to the filtration cartridge. Cap and vortex vigorously (~1 min). Alternatively, sonicate (&#62;10 min) to disperse the protein pellet.
		<ol>
			<li>Heat the sample at 95 &#176;C for 5 min). Repeat the mixing step after heating. 
			NOTE: Laemmli gel loading buffer can re-solubilize the protein pellet. However, SDS-containing samples are incompatible with trypsin digestion and reversed-phase LC and MS.</li>
		</ol>
		</li>
		<li>(Option 2) Prepare a solution of 80% (v/v) formic acid in water. Prechill the acid solution (-20 &#176;C), as well as the filtration cartridge containing precipitated protein.
		<ol>
			<li>Dispense 50 &#181;L of cold formic acid into the cartridge; cap and vortex for 30 s. Return to the freezer (-20 &#176;C) for 10 min.</li>
			<li>Vortex the cartridge again for 30 s. Then, repeat the chilling and mixing cycle one more time (10 min, -20 &#176;C, 30 s vortex).</li>
			<li>Add water to a final 500 &#181;L, diluting the formic acid to 8%. 
			NOTE: The cold formic acid protocol is incompatible with subsequent trypsin digestion but is compatible with LC-MS.</li>
		</ol>
		</li>
		<li>(Option 3) Add 50 &#181;L of freshly prepared 8 M urea in water to the filtration cartridge. Sonicate for 30 min.
		<ol>
			<li>Allow the cartridge to incubate on the benchtop for 1 h (up to overnight).</li>
			<li>Dilute the 8 M urea a minimum 5-fold with water or appropriate buffer. 
			NOTE: Once diluted, the urea solubilization protocol is compatible with subsequent trypsin digestion, as well as LC-MS.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
7. Protein digestion
<ol>
	<li>For bottom-up MS analysis, subject the re-solubilized proteins to enzymatic digestion using one of the two methods mentioned below.
	<ol>
		<li>(Option 1) For formic acid resolubilization, reduce the initial volume of 80% formic acid in step 6.2.2.1 to 25 &#181;L. In step 6.2.2.3, use 375 &#181;L of water to dilute the formic acid to 5% (v/v).</li>
		<li>Dispense pepsin into the cartridge at an approximate protein to enzyme ratio of 50:1. With a plug attached to the filtration cartridge, incubate the sample overnight at room temperature.</li>
	</ol>
	</li>
	<li>(Option 2) For resolubilization in urea, ensure a pH between 8 and 8.3 with the inclusion of 100 mM of Tris or ammonium bicarbonate in step 6.2.3.2.
	<ol>
		<li>Add trypsin at an approximate protein to enzyme mass ratio of 50:1. With a plug attached to the cartridge, incubate the sample overnight in a warm water bath at 37 &#176;C.</li>
		<li>Terminate the digestion by acidifying the solution with 10% TFA to a final 1%.</li>
	</ol>
	</li>
	<li>Recover the pepsin- or trypsin-digested protein by removing the plug from the base of the filter and centrifuging the cartridge contained within a clean vial (2 min, 5000 x g, room temperature).</li>
</ol>
8. SPE clean-up
NOTE: For additional sample desalting following digestion or solvent exchange, the sample can be subject to reversed-phase clean-up as described.
<ol>
	<li>Prime an SPE cartridge (see Table of Materials) by passing 300 &#181;L of methanol (2 min, 400 x g) followed by 300 &#181;L of 5% acetonitrile/0.1% of TFA (2 min, 400 x g).</li>
	<li>Connect the primed SPE cartridge to the base of the filtration cartridge containing re-solubilized or digested protein.</li>
	<li>Spin the protein through the SPE cartridge (5 min, 800 x g at room temperature). If solvent remains in the upper cartridge, return the cartridge to the centrifuge and repeat the spin. 
	NOTE: Passing the sample through the SPE cartridge a second time may improve recovery.</li>
	<li>Add 300 &#181;L of 5% acetonitrile/0.1% of TFA in water to the cartridge. To wash, flow through the SPE cartridge (2 min, 2000 x g). Discard the flow-through.</li>
	<li>For LMW proteins or digested peptides, elute the sample by flowing 300 &#181;L of 50% acetonitrile/0.1%&#160;TFA (5 min, 2500 x g).</li>
	<li>For intact proteins, follow step 8.5 with an additional elution step using 300 &#181;L of 75% acetonitrile/0.1% TFA. Combine the two resulting extracts. 
	NOTE: Step 8.6 is optional.</li>
</ol>

The Doucette laboratory conceived of and patented the ProTrap XG employed in this study. AAD is also a founding partner of Proteoform Scientific, which commercialized the sample preparation cartridge.

Optimal MS characterization is achieved when residual SDS is depleted below 10 ppm. While alternative approaches, such as FASP and on-bead digestion, offer quantitative SDS depletion with variable recovery31,32,33, the primary objective of precipitation is to maximize purity and yield simultaneously. This depends on effectively isolating the supernatant (containing the SDS) without disturbing the protein pellet. With vial-based ...

Proteome analysis by mass spectrometry has become increasingly rigorous, owing to the enhanced sensitivity, resolution, scan speed, and versatility of modern MS instruments. MS advances contribute to greater protein identification efficiency and more precise quantitation1,2,3,4,5. With improved MS instrumentation, researchers demand a correspondingly consistent front-end sample preparation strategy capable of quantitative recovery of high-purity proteins in minimal time across all stages of the workflow6,7,8,9,10,11. To accurately reflect the proteome status of a biological system, proteins must be isolated from the native sample matrix in an efficient and unbiased fashion. To this end, including a denaturing surfactant, such as sodium dodecyl sulfate (SDS), ensures efficient protein extraction and solubilization12. However, SDS strongly interferes with electrospray ionization, causing severe MS signal suppression if not properly eliminated13.
Various SDS depletion strategies are available for subsequent proteome analysis, such as the retention of proteins above a molecular weight cutoff filter contained within disposable spin cartridges14,15,16. The filter-aided sample preparation method (FASP) is favored as it effectively depletes SDS below 10 ppm, facilitating optimal MS. However, protein recovery with FASP is variable, which prompted the exploration of other techniques. Chromatographic approaches that selectively capture protein (or surfactant) have evolved into various convenient cartridges or bead-based formats17,18,19,20,21. Given these simple and (ideally) consistent strategies to protein purification, the classical approach of protein precipitation with organic solvents is often overlooked as a promising approach to protein isolation. While solvent precipitation is shown to deplete SDS below critical levels successfully, protein recovery has been a longstanding concern of this approach. Multiple groups have observed a protein recovery bias, with unacceptably low precipitation yields as a function of protein concentration, molecular weight, and hydrophobicity22,23. Due to the diversity of precipitation protocols reported in the literature, standardized precipitation conditions were developed. In 2013, Crowell et al. first reported the dependence of ionic strength on the precipitation efficiency of proteins in 80% acetone24. For all proteins examined, the addition of up to 30 mM sodium chloride was shown to be essential to maximize yields (up to 100% recovery). More recently, Nickerson et al. showed that the combination of even higher ionic strength (up to 100 mM) with elevated temperature (20 &#176;C) during acetone precipitation gave near quantitative recovery in 2-5 min25. A slight drop in the recovery of low molecular weight (LMW) proteins was observed. Therefore, a subsequent report by Baghalabadi et al. demonstrated the successful recovery of LMW proteins and peptides (&#8804;5 kDa) by combining specific salts, particularly zinc sulfate, with a higher level of organic solvent (97% acetone)26.
While refining the precipitation protocol lends a more reliable protein purification strategy for MS-based proteomics, the success of conventional precipitation relies heavily on user technique. A primary goal of this work is to present a robust precipitation strategy that facilitates the isolation of the protein pellet from the contaminating supernatant. A disposable filtration cartridge was developed to eliminate pipetting by isolating aggregated protein above a porous PTFE membrane filter27. MS-interfering components in the supernatant are effectively removed in a short, low-speed&#160;centrifugation step. The disposable filter cartridge also offers an interchangeable SPE cartridge, which facilitates subsequent sample clean-up following resolubilization and optional protein digestion, ahead of mass spectrometry.
A series of recommended proteome precipitation workflows are presented here, including modified acetone and chloroform/methanol/water28 protocols, in a conventional (vial-based) and a semi-automated format in a disposable two-state filtration and extraction cartridge. The resulting protein recoveries and SDS depletion efficiencies are highlighted, together with bottom-up LC-MS/MS proteome coverage, to demonstrate the expected outcome from each protocol. The practical benefits and drawbacks associated with each approach are discussed.

<table><tbody><tr><td>Acetone</td><td>Fisher Scientific</td><td>AC177170010</td><td>&amp;le;0.002 % aldehyde</td></tr><tr><td>Acetonitrile</td><td>Fisher Scientific</td><td>A998-4</td><td>HPLC grade</td></tr><tr><td>Ammonium Bicarbonate</td><td>Millipore Sigma</td><td>A6141-1KG</td><td>solid</td></tr><tr><td>Beta mercaptoethanol</td><td>Millipore Sigma</td><td>M3148-25ML</td><td>Molecular biology grade</td></tr><tr><td>Bromophenol blue</td><td>Millipore Sigma</td><td>B8026-5G</td><td>Bromophenol blue sodium salt</td></tr><tr><td>Chloroform</td><td>Fisher Scientific</td><td>C298-400</td><td>Chloroform</td></tr><tr><td>Formic Acid</td><td>Honeywell</td><td>56302</td><td>Eluent additive for LC-MS</td></tr><tr><td>Fusion Lumos Mass Spectrometer</td><td>ThermoFisher Scientific</td><td></td><td>for analysis of standard protein mixture</td></tr><tr><td>Glycerol</td><td>Millipore Sigma</td><td>356352-1L-M</td><td>For molecular biology, &gt; 99%</td></tr><tr><td>Isopropanol</td><td>Fisher Scientific</td><td>A4641</td><td>HPLC grade</td></tr><tr><td>Methanol</td><td>Fisher Scientific</td><td>A452SK-4</td><td>HPLC grade</td></tr><tr><td>Microcentrifuge</td><td>Fisher Scientific</td><td>75-400-102</td><td>up to 21,000 xg</td></tr><tr><td>Microcentrifuge Tube (1.5 mL)</td><td>Fisher Scientific</td><td>05-408-130</td><td>tapered bottom</td></tr><tr><td>Microcentrifuge Tube&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (2 mL)</td><td>Fisher Scientific</td><td>02-681-321</td><td>rounded bottom</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (0.1-10 &amp;mu;L)</td><td>Fisher Scientific</td><td>21-197-28</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (1-200 &amp;mu;L)</td><td>Fisher Scientific</td><td>07-200-302</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipette Tips&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; (200-1000 &amp;mu;L)</td><td>Fisher Scientific</td><td>07-200-303</td><td>Universal pipet tip, non-sterile</td></tr><tr><td>Micropipettes</td><td>Fisher Scientific</td><td>13-710-903</td><td>Micropipet Trio pack</td></tr><tr><td>Pepsin</td><td>Millipore Sigma</td><td>P0525000</td><td>Lyophilized powder,&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp;&amp;nbsp; &gt;3200 units/ mg</td></tr><tr><td>ProTrap XG</td><td>Proteoform Scientific</td><td>PXG-0002</td><td>50 complete units per box</td></tr><tr><td>Sodium Chloride</td><td>Millipore Sigma</td><td>S9888-1KG</td><td>ACS reagent, &gt;99 %</td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>ThermoFisher Scientific</td><td>28312</td><td>powdered solid</td></tr><tr><td>timsTOF Pro Mass Spectrometer</td><td>Bruker</td><td></td><td>for analysis of liver proteome extract</td></tr><tr><td>Trifluoroacetic Acid</td><td>ThermoFisher Scientific</td><td>L06374.AP</td><td>99%</td></tr><tr><td>Tris</td><td>Fisher Scientific</td><td>BP152-500</td><td>Molecular biology grade</td></tr><tr><td>Trypsin</td><td>Millipore Sigma</td><td>9002-07-7</td><td>From bovine pancreas, TPCK-treated</td></tr><tr><td>Urea</td><td>Bio-Rad</td><td>1610731</td><td>solid</td></tr><tr><td>Water (deionized)</td><td>Sartorius Arium Mini Water Purification System</td><td>76307-662</td><td>Type 1 ultrapure (18.2 M&amp;Omega; cm)</td></tr><tr><td>Zinc Sulfate</td><td>Millipore Sigma</td><td>307491-100G</td><td>solid</td></tr></tbody></table>

organic solvent-based protein precipitation for robust proteome purification ahead of mass spectrometry

Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end approaches to isolate, enrich, and process proteins ahead of MS are critical for successful proteome characterization. Low, inconsistent protein recovery and residual impurities such as surfactants are detrimental to MS analysis. Protein precipitation is often considered unreliable, time-consuming, and technically challenging to perform compared to other sample preparation strategies. These concerns are overcome by employing optimal protein precipitation protocols. For acetone precipitation, the combination of specific salts, temperature control, solvent composition, and precipitation time is critical, while the efficiency of chloroform/methanol/water precipitation depends on proper pipetting and vial manipulation. Alternatively, these precipitation protocols are streamlined and semi-automated within a disposable spin cartridge. The expected outcomes of solvent-based protein precipitation in the conventional format and using a disposable, two-stage filtration and extraction cartridge are illustrated in this work. This includes the detailed characterization of proteomic mixtures by bottom-up LC-MS/MS analysis. The superior performance of SDS-based workflows is also demonstrated relative to non-contaminated protein.

Figure 4 summarizes the expected SDS depletion following vial-based or cartridge-facilitated precipitation of proteins in a disposable filter cartridge using acetone. Conventional overnight incubation (-20 °C) in acetone is compared to the rapid acetone precipitation protocol at room temperature (step 2), as well as CMW precipitation (step 4). Residual SDS was quantified by the methylene blue active substances (MBAS) assay29. Briefly, 100 µL sample was combined with 1...

The present protocol describes solvent-based protein precipitation under controlled conditions for robust and rapid recovery and purification of proteome samples prior to mass spectrometry.

While multiple advances in mass spectrometry (MS) instruments have improved qualitative and quantitative proteome analysis, more reliable front-end ...

These protocols are designed to maximize the recovery as well as the purity of proteins through solvent-based precipitation. We're gonna show you a couple tricks to perform precipitation in vials, as well as a semi-automated approach in a spin cartridge format. We have also optimized the precipitation protocol to be consistent and quick.The whole protocol only takes a few minutes. These protocols demonstrate that solvent precipitation of proteins is ideal for sample prep ahead of mass spectrometry. The protocols will fit seamlessly into bottom up as well as top down pro workflows.Demonstrating for us today will be Ziheng Dang and Victoria Miller, Senior scientist at Proteoform Scientific in Halifax, Nova Scotia Pipette 90 microliters of particulate free protein solution into a polypropylene micro centrifuge tube. Then add 10 microliters of one mole aqueous sodium chloride and 400 microliters of acetone into the sample solution, cap and tap the vial gently to combine the solvents. Allow the vial to incubate it.Room temperature undisturbed for a minimum of two minutes following incubation, centrifuge the samples noting the orientation of the vial and spin for a minimum of two minutes at 10, 000 RCF or higher at room temperature. After the centrifugation, a visible white pellet is observed at the bottom of the tube. At this point, uncap the tube and decant the supernate into a waste container by inverting the tube.Use a paper towel to draw any residual solvent from the vial. For SDS containing samples. Carefully dispense 400 microliters of fresh acetone without disturbing the pellet.Immediately centrafuse the sample for one minute at 10, 000 times G or higher at room temperature by placing the vial into the rotor in the same orientation as the initial spin. Decount the wash solvent as described previously. Dry the sample with the cap open for approximately one minute.In a fume hood, before a pellet solubilization. Dispense 100 microliters of the protein solution into a polypropylene vial. In the fume hood, add a 400 microliters of methanol followed by 100 microliters of chloroform.Cap the vial and vortex briefly to mix. Quickly dispense 300 microliters of water directly into the center of the vial. Cap the vial and allow it to sit on the bench top undisturbed for one minute, after place the polypropylene vial in a centrifuge and spin it for five minutes at 10, 000 times G or higher at room temperature.The vial is observed to have two solvent layers and a visible white pellet til the vial at approximately 45 degrees. Remove approximately 700 microliters of the solvent from the upper layer at a uniform rate using a large one milliliter micro pipette tip initially and later with a 200 microliter pipette tip until it forms a bead in the vial. Add 400 microliters of fresh methanol to the sample vial without disturbing the pellet by dispensing the solvent down the side of the vial.Cap the vial and combine the solvent layers by gently rocking the vial to swirl the solvents together. To observe the white protein pellet noting the orientation of the vial in the rotor centrifuge for a minimum of 10 minutes at 10, 000 times G or higher at room temperature. Angle the vial with the pellet facing down.Place a pipette tip along the upper edge of the vial and remove the supernatant with a one milliliter micro pipette tip at a slow but uniform rate. Retaining approximately 20 microliters of solvent. Allow the residual solvent to dry in the fume hood.To precipitate the protein. Using a disposable filtration cartridge attach the plug to the upper filtration cartridge. Combine 90 microliters of protein solution 10 microliters of one mole aquas, sodium chloride and 400 microliters of acetone.Incubate for a minimum of two minutes on the bench top, centrifuge the prepared protein sample with the plug still attached to the filtration cartridge for two minutes at 2, 500 times G.At room temperature, invert the cartridge and unscrew and remove the plug from the cartridge base. Place the filtration cartridge in a clean vial centrifuge the protein sample for three minutes at 500 times G at room temperature. Discard the flow through solvent from the lower vial.Wash the protein pellet by adding 400 microliters of acetone to the filtration cartridge, centrifuge for three minutes at 500 times G at room temperature or until no solvent remains in the upper cartridge. Following the previously demonstrated protein precipitation protocols. Re solubilize the protein by wetting the membrane at the base of filtration cartridge by dispensing two to five microliters of ISO propanol directly to the membrane immediately before proceeding to the re solubilization protocol for re solubilization of the protein pellet prepare 80%volume by volume solution of formic acid in water.Pre chill this acid solution at minus 20 degrees Celsius and the filtration cartridge containing precipitated protein. Dispense 50 microliters of cold diluted formic acid into the chilled cartridge cap and vortex for 30 seconds. Return the filtration cartridge to the freezer at minus 20 degrees Celsius for 10 minutes.Repeat the previous vortex and incubation steps. Add 450 microliters of water to a final volume of 500 microliters. Diluting the formic acid to 8%Invert the cartridge unscrew and remove the plug from the cartridge base and attach the SPE cartridge to the vial.Spin the protein through the SPE cartridge in a centrifuge at room temperature for five minutes at 800 times G.If solvent remains in the upper cartridge return the cartridge to the centrifuge and repeat the spin. Rinse by adding 300 microliters of 5%acetonitrile with 0.1%of TFA in water to the SPE cartridge, flow through the SPE cartridge for two minutes at 2000 times G.For low molecular weight proteins or digested peptides elute the sample by flowing 300 microliters. A 50%acetonitrile with 0.1%of TFA for five minutes at 2, 500 times G.For intact proteins.Follow up with an additional elusion step using 300 microliters of 75%acetonitrile with 0.1%Trifluoro ascetic acid. Combine the two resulting extracts. The figure compares the SDS depletion following vial based or precipitation of proteins in a disposable filter cartridge.Using acetone with the conventional rapid and CMW protocols. All approaches reduce SDS to permit optimal mass spectrometry analysis. This figure shows quantitative protein recovery by following rapid acetone precipitation and CMW precipitation through SDS page analysis of a processed yeast total cell lysate.Consistent recovery was obtained with all precipitation protocols. Precipitation in a disposable filtration cartridge eliminates the need to carefully pipette SDS containing supernatant while retaining the aggregated proteins above a membrane filter. Thus no visible bands were detected in the supernatant fractions across three independent replicates.The cartridge based protein precipitation demonstrates superior recovery of a Pepsin digested sample of bovine plasma relative to vial based precipitation. More significant differences in yield are noted in the cartridge when employing sodium chloride while employing zinc solvent resulted in the highest yield. This figure quantifies the peptide peak intensity from each peptide sample with a ratio above one reflecting a higher peptide abundance for samples processed in the disposable filter cartridges.The figure compares the number of identified peptides per protein across the three replicate sample preparations. The correlation coefficients of 0.94 to 0.95 in these graphs demonstrate the high consistency of the sample preparation approach for bottom up mass spectrometry analysis. So once that pellet is formed you really wanna avoid disturbing it.If you keep it as an intact pellet you'll get a higher yield. So make sure not to forget to wash that pellet with extra acetone. This is making sure that we're getting rid of the SDS keeping our sample as clean as possible and that gives us the best mass spectrometry analysis.So we found that removing the excess solvent by inverting the is a lot simpler than trying to pipette the sample. You'll get more consistent yields this way. Performing acetone precipitation at room temperature resulted in more consistent and faster recovery than traditional precipitation in the freezer.

organic-solvent-based-protein-precipitation-for-robust-proteome

Research

JoVE Journal

Chemistry

Digital Microfluidics is a technique characterized by the manipulation of discrete droplets (~nL - mL) on an array of electrodes by the application of electrical fields. It is well-suited for carrying out rapid, sequential, miniaturized automated biochemical assays. Here, we report a platform capable of automating several proteomic processing steps.

Digital Microfluidics for Automated Proteomic Processing

We present a variation of the QUICK (QUantitative Immunoprecipitation Combined with Knockdown) approach that was introduced previously to distinguish between true and false protein-protein interactions. Our approach is based on 15N metabolic labeling, the modulation of affinities of protein-protein interactions by the presence/absence of ATP, immunoprecipitation, and quantitative mass spectrometry.

A Protocol for the Identification of Protein-protein Interactions Based on 15N Metabolic Labeling, Immunoprecipitation, Quantitative Mass Spectrometry and Affinity Modulation

A common protein extraction protocol using urea/thiourea/SDS buffer for human and mice brain tissue allows indentification of proteins by 2D-DIGE and their subsequent characterization by mini 2DE immunoblotting. This method enables one to obtain more reproducible and reliable results from human biopsies and experimental models.

Consensus Brain-derived Protein, Extraction Protocol for the Study of Human and Murine Brain Proteome Using Both 2D-DIGE and Mini 2DE Immunoblotting

Stable isotope labeling of peptides by reductive dimethylation (ReDi labeling) is a rapid, inexpensive strategy for accurate mass spectrometry-based quantitative proteomics. Here we demonstrate a robust method for preparation and analysis of protein mixtures using the ReDi approach that can be applied to nearly any sample type.

Quantitative Proteomics Using Reductive Dimethylation for Stable Isotope Labeling

The reliability of results in metabolomics experiments depends on the effectiveness and reproducibility of the sample preparation. Described is a rigorous and in-depth method that enables extraction of metabolites from biological fluids with the option of subsequently analyzing up to thousands of compounds, or just the compound classes of interest.

Multi-step Preparation Technique to Recover Multiple Metabolite Compound Classes for In-depth and Informative Metabolomic Analysis

This protocol is for the extraction and concentration of protein and DNA from microbial biomass collected from seawater, followed by the generation of tryptic peptides suitable for tandem mass spectrometry-based proteomic analysis.

An Aquatic Microbial Metaproteomics Workflow: From Cells to Tryptic Peptides Suitable for Tandem Mass Spectrometry-based Analysis

Protein-protein and protein-metabolite interactions are crucial for all cellular functions. Herein, we describe a protocol that allows parallel analysis of these interactions with a protein of choice. Our protocol was optimized for plant cell cultures and combines affinity purification with mass spectrometry-based protein and metabolite detection.

2 in 1: One-step Affinity Purification for the Parallel Analysis of Protein-Protein and Protein-Metabolite Complexes

Described is a proteomics workflow for identifying protein interaction partners from a nuclear subcellular fraction using immunoaffinity enrichment of a given protein of interest and label-free mass spectrometry. The workflow includes subcellular fractionation, immunoprecipitation, filter aided sample preparation, offline cleanup, mass spectrometry, and a downstream bioinformatics pipeline.

Label-Free Immunoprecipitation Mass Spectrometry Workflow for Large-scale Nuclear Interactome Profiling

The present protocol describes the extraction of non-protein amino acids from biological matrices via trichloroacetic acid (TCA) protein precipitation and acid hydrolysis before analysis using liquid chromatography-tandem mass spectrometry.

Extraction of Non-Protein Amino Acids from Cyanobacteria for Liquid Chromatography-Tandem Mass Spectrometry Analysis

This protocol describes a method for collecting tear samples using Schirmer strips and an integrated quantitative workflow for discovering non-invasive tear protein biomarkers. The suspension trapping sample preparation workflow enables fast and robust tear sample preparation and mass spectrometric analysis, resulting in higher peptide recovery yields and protein identification than standard in-solution procedures.

A Protein Suspension-Trapping Sample Preparation for Tear Proteomics by Liquid Chromatography-Tandem Mass Spectrometry

Here, we establish a mass spectrometry-based proteomic method using isolated regions of interest in formalin-fixed, paraffin-embedded tissue sections. This protocol is used to analyze proteome from specific tissue areas in archived formalin-fixed, paraffin-embedded tissue sections.

A Streamlined Approach for Mass Spectrometry-Based Proteomics Using Selected Tissue Regions

Learn organic solvent-based protein precipitation for robust proteome purification ahead of mass spectrometry. This optimized protocol maximizes recovery and purity in minutes for bottom-up and top-down workflows.

Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry

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