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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Acetone</td>
			<td >Fisher Scientific</td>
			<td >AC177170010</td>
			<td >&#8804;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, &#62; 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&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; (2 mL)</td>
			<td >Fisher Scientific</td>
			<td >02-681-321</td>
			<td >rounded bottom</td>
		</tr>
		<tr >
			<td >Micropipette Tips&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; (0.1-10 &#956;L)</td>
			<td >Fisher Scientific</td>
			<td >21-197-28</td>
			<td >Universal pipet tip, non-sterile</td>
		</tr>
		<tr >
			<td >Micropipette Tips&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; (1-200 &#956;L)</td>
			<td >Fisher Scientific</td>
			<td >07-200-302</td>
			<td >Universal pipet tip, non-sterile</td>
		</tr>
		<tr >
			<td >Micropipette Tips&#160;&#160;&#160;&#160;&#160;&#160;&#160; (200-1000 &#956;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,&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; &#62;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, &#62;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&#937; 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

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 &#176;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 &#181;L sample was combi...

Watch this Scientific Journal Video about Organic Solvent-Based Protein Precipitation for Robust Proteome Purification Ahead of Mass Spectrometry at JoVE.com

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

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 ...

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

Microsoft Bing

Research

Education

JoVE Core

JoVE Journal

Analytical Chemistry

Chemistry

Unit 1

Complexometric Titration, Precipitation Titration, and Gravimetry

SCIENTISTS IN ACTION

9.5K Views.  Dalhousie University. 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.

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

Analyzing Protein Dynamics Using Hydrogen Exchange Mass Spectrometry

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique amino acid sequence, it is only realized by the folding of the polypeptide chain into a single defined three-dimensional arrangement or more commonly into an ensemble of interconverting conformations. Investigating the connection between protein conformation and its function is therefore essential for a complete understanding of how proteins are able to fulfill their great variety of tasks. One possibility to study conformational changes a protein undergoes while progressing through its functional cycle is hydrogen-1H/2H-exchange in combination with high-resolution mass spectrometry (HX-MS). HX-MS is a versatile and robust method that adds a new dimension to structural information obtained by e.g. crystallography. It is used to study protein folding and unfolding, binding of small molecule ligands, protein-protein interactions, conformational changes linked to enzyme catalysis, and allostery. In addition, HX-MS is often used when the amount of protein is very limited or crystallization of the protein is not feasible. Here we provide a general protocol for studying protein dynamics with HX-MS and describe as an example how to reveal the interaction interface of two proteins in a complex. &#160;&#160;

Protein conformation and dynamics are key to understanding the relationship between protein structure and function. Hydrogen exchange coupled with high-resolution mass spectrometry is a versatile method to study the conformational dynamics of proteins as well as characterizing protein-ligand and protein-protein interactions, including contact interfaces and allosteric effects.

All cellular processes depend on the functionality of proteins. Although the functionality of a given protein is the direct consequence of its unique ...

Analysis of Volatile and Oxidation Sensitive Compounds Using a Cold Inlet System and Electron Impact Mass Spectrometry

This video presents a protocol for the mass spectrometrical analysis of volatile and oxidation sensitive compounds using electron impact ionization. The analysis of volatile and oxidation sensitive compounds by mass spectrometry is not easily achieved, as all state-of-the-art mass spectrometric methods require at least one sample preparation step, e.g., dissolution and dilution of the analyte (electrospray ionization), co-crystallization of the analyte with a matrix compound (matrix-assisted laser desorption/ionization), or transfer of the prepared samples into the ionization source of the mass spectrometer, to be conducted under atmospheric conditions. Here, the use of a sample inlet system is described which enables the analysis of volatile metal organyls, silanes, and phosphanes using a sector field mass spectrometer equipped with an electron impact ionization source. All sample preparation steps and the sample introduction into the ion source of the mass spectrometer take place either under air-free conditions or under vacuum, enabling the analysis of compounds highly susceptible to oxidation. The presented technique is especially of interest for inorganic chemists, working with metal organyls, silanes, or phosphanes, which have to be handled using inert conditions, such as the Schlenk technique. The principle of operation is presented in this video.

This video presents a protocol for the mass spectrometrical analysis of volatile and oxidation sensitive compounds using electron impact ionization. The presented technique is especially of interest for inorganic chemists, working with metal organyls, silanes, or phosphanes which have to be handled using inert conditions, such as the Schlenk technique.

This video presents a protocol for the mass spectrometrical analysis of volatile and oxidation sensitive compounds using electron impact ionization. The ...

An HS-MRM Assay for the Quantification of Host-cell Proteins in Protein Biopharmaceuticals by Liquid Chromatography Ion Mobility QTOF Mass Spectrometry

The analysis of low-level (1-100 ppm) protein impurities (e.g., host-cell proteins (HCPs)) in protein biotherapeutics is a challenging assay requiring high sensitivity and a wide dynamic range. Mass spectrometry-based quantification assays for proteins typically involve protein digestion followed by the selective reaction monitoring/multiple reaction monitoring (SRM/MRM) quantification of peptides using a low-resolution (Rs ~1,000) tandem quadrupole mass spectrometer. One of the limitations of this approach is the interference phenomenon observed when the peptide of interest has the "same" precursor and fragment mass (in terms of m/z values) as other co-eluting peptides present in the sample (within a 1-Da window). To avoid this phenomenon, we propose an alternative mass spectrometric approach, a high selectivity (HS) MRM assay that combines the ion mobility separation of peptide precursors with the high-resolution (Rs ~30,000) MS detection of peptide fragments. We explored the capabilities of this approach to quantify low-abundance peptide standards spiked in a monoclonal antibody (mAb) digest and demonstrated that it has the sensitivity and dynamic range (at least 3 orders of magnitude) typically achieved in HCP analysis. All six peptide standards were detected at concentrations as low as 0.1 nM (1 femtomole loaded on a 2.1-mm ID chromatographic column) in the presence of a high-abundance peptide background (2 &#181;g of a mAb digest loaded on-column). When considering the MW of rabbit phosphorylase (97.2 kDa), from which the spiked peptides were derived, the LOQ of this assay is lower than 50 ppm. Relative standard deviations (RSD) of peak areas (n = 4 replicates) were less than 15% across the entire concentration range investigated (0.1-100 nM or 1-1,000 ppm) in this study.

Here, we describe a chromatographic assay coupled with the ion mobility separation of peptide precursors followed by the high-resolution (~30,000) MS-detection of peptide fragments for the quantification of spiked peptide standards in a monoclonal antibody digest.

The analysis of low-level (1-100 ppm) protein impurities (e.g., host-cell proteins (HCPs)) in protein biotherapeutics is a challenging assay requiring ...

Fizzy Extraction of Volatile Organic Compounds Combined with Atmospheric Pressure Chemical Ionization Quadrupole Mass Spectrometry

Chemical analysis of volatile and semivolatile compounds dissolved in liquid samples can be challenging. The dissolved components need to be brought to the gas phase, and efficiently transferred to a detection system. Fizzy extraction takes advantage of the effervescence phenomenon. First, a carrier gas (here, carbon dioxide) is dissolved in the sample by applying overpressure and stirring the sample. Second, the sample chamber is decompressed abruptly. Decompression leads to the formation of numerous carrier gas bubbles in the sample liquid. These bubbles assist the release of the dissolved analyte species from the liquid to the gas phase. The released analytes are immediately transferred to the atmospheric pressure chemical ionization interface of a triple quadrupole mass spectrometer. The ionizable analyte species give rise to mass spectrometric signals in the time domain. Because the release of the analyte species occurs over short periods of time (a few seconds), the temporal signals have high amplitudes and high signal-to-noise ratios. The amplitudes and areas of the temporal peaks can then be correlated with concentrations of the analytes in the liquid samples subjected to fizzy extraction, which enables quantitative analysis. The advantages of fizzy extraction include: simplicity, speed, and limited use of chemicals (solvents).

Fizzy extraction is a new laboratory technique for analysis of volatile and semivolatile compounds. A carrier gas is dissolved in the liquid sample by applying overpressure and stirring the sample. The sample chamber is then decompressed. The analyte species are liberated to the gas phase due to effervescence.

Chemical analysis of volatile and semivolatile compounds dissolved in liquid samples can be challenging. The dissolved components need to be brought to ...

Real-time Breath Analysis by Using Secondary Nanoelectrospray Ionization Coupled to High Resolution Mass Spectrometry

Exhaled volatile organic compounds (VOCs) have aroused considerable interest, since they can serve as biomarkers for disease diagnosis and environmental exposure in a non-invasive manner. In this work, we present a protocol to characterize the exhaled VOCs in real time by using secondary nanoelectrospray ionization coupled to high resolution mass spectrometry (Sec-nanoESI-HRMS). The homemade Sec-nanoESI source was readily set up based on a commercial nanoESI source. Hundreds of peaks were observed in the background-subtracted mass spectra of exhaled breath, and the mass accuracy values are -4.0-13.5 ppm and -20.3-1.3 ppm in the positive and negative ion detection modes, respectively. The peaks were assigned with accurate elemental composition according to the accurate mass and isotopic pattern. Less than 30 s is used for one exhalation measurement, and it takes approximately 7 min for six replicated measurements.

A protocol for characterizing chemical composition of exhaled breath in real time by using secondary nanoelectrospray ionization coupled to high resolution mass spectrometry is demonstrated.

Exhaled volatile organic compounds (VOCs) have aroused considerable interest, since they can serve as biomarkers for disease diagnosis and environmental ...

Synthesis and Mass Spectrometry Analysis of Oligo-peptoids

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have been thought of as a type of molecular information storage. Mass spectrometry analysis has been considered the method of choice for sequencing peptoids. Peptoids can be synthesized via solid phase chemistry using a repeating two-step reaction cycle. Here we present a method to manually synthesize oligo-peptoids and to analyze the sequence of the peptoids using tandem mass spectrometry (MS/MS) techniques. The sample peptoid is a nonamer consisting of alternating N-(2-methyloxyethyl)glycine (Nme) and N-(2-phenylethyl)glycine (Npe), as well as an N-(2-aminoethyl)glycine (Nae) at the N-terminus. The sequence formula of the peptoid is Ac-Nae-(Npe-Nme)4-NH2, where Ac is the acetyl group. The synthesis takes place in a commercially available solid-phase reaction vessel. The rink amide resin is used as the solid support to yield the peptoid with an amide group at the C-terminus. The resulting peptoid product is subjected to sequence analysis using a triple-quadrupole mass spectrometer coupled to an electrospray ionization source. The MS/MS measurement produces a spectrum of fragment ions resulting from the dissociation of charged peptoid. The fragment ions are sorted out based on the values of their mass-to-charge ratio (m/z). The m/z values of the fragment ions are compared against the nominal masses of theoretically predicted fragment ions, according to the scheme of peptoid fragmentation. The analysis generates a fragmentation pattern of the charged peptoid. The fragmentation pattern is correlated to the monomer sequence of the neutral peptoid. In this regard, MS analysis reads out the sequence information of the peptoids.

A protocol is described for the manual synthesis of oligo-peptoids followed by sequence analysis by mass spectrometry.

Peptoids are sequence-controlled peptide-mimicking oligomers consisting of N-alkylated glycine units. Among many potential applications, peptoids have ...

Analyzing Protein Architectures and Protein-Ligand Complexes by Integrative Structural Mass Spectrometry

Proteins are an important class of biological macromolecules that play many key roles in cellular functions including gene expression, catalyzing metabolic reactions, DNA repair and replication. Therefore, a detailed understanding of these processes provides critical information on how cells function. Integrative structural MS methods offer structural and dynamical information on protein complex assembly, complex connectivity, subunit stoichiometry, protein oligomerization and ligand binding. Recent advances in integrative structural MS have allowed for the characterization of challenging biological systems including large DNA binding proteins and membrane proteins. This protocol describes how to integrate diverse MS data such as native MS and ion mobility-mass spectrometry (IM-MS) with molecular dynamics simulations to gain insights into a helicase-nuclease DNA repair protein complex. The resulting approach provides a framework for detailed studies of ligand binding to other protein complexes involved in important biological processes.

Mass spectrometry (MS) has emerged as an important tool for the investigation of structure and dynamics of macromolecular assemblies. Here, we integrate MS-based approaches to interrogate protein complex formation and ligand binding.

Proteins are an important class of biological macromolecules that play many key roles in cellular functions including gene expression, catalyzing ...

Mass Spectrometry-Guided Genome Mining as a Tool to Uncover Novel Natural Products

The chemical space covered by natural products is immense and widely unrecognized. Therefore, convenient methodologies to perform wide-ranging evaluation of their functions in nature and potential human benefits (e.g., for drug discovery applications) are desired. This protocol describes the combination of genome mining (GM) and molecular networking (MN), two contemporary approaches that match gene cluster-encoded annotations in whole genome sequencing with chemical structure signatures from crude metabolic extracts. This is the first step towards the discovery of new natural entities. These concepts, when applied together, are defined here as MS-guided genome mining. In this method, the main components are previously designated (using MN), and structurally related new candidates are associated with genome sequence annotations (using GM). Combining GM and MN is a profitable strategy to target new molecule backbones or harvest metabolic profiles in order to identify analogues from already known compounds.

A mass spectrometry-guided genome mining protocol is established and described here. It is based on genome sequence information and LC-MS/MS analysis and aims to facilitate identification of molecules from complex microbial and plant extracts.

The chemical space covered by natural products is immense and widely unrecognized. Therefore, convenient methodologies to perform wide-ranging evaluation ...

Capillary Electrophoresis Mass Spectrometry Approaches for Characterization of the Protein and Metabolite Corona Acquired by Nanomaterials

The adsorption of biomolecules from surrounding biological matrices to the surface of nanomaterials (NMs) to form the corona has been of interest for the past decade. Interest in the bio-nano interface arises from the fact that the biomolecular corona confers a biological identity to NMs and thus causes the body to identify them as &#8220;self&#8221;. For example, previous studies have demonstrated that the proteins in the corona are capable of interacting with membrane receptors to influence cellular uptake and established that the corona is responsible for cellular trafficking of NMs and their eventual toxicity. To date, most research has focused upon the protein corona and overlooked the possible impacts of the metabolites included in the corona or synergistic effects between components in the complete biomolecular corona. As such, this work demonstrates methodologies to characterize both the protein and metabolite components of the biomolecular corona using bottom-up proteomics and metabolomics approaches in parallel. This includes an on-particle digest of the protein corona with a surfactant used to increase protein recovery, and a passive characterization of the metabolite corona by analyzing metabolite matrices before and after NM exposures. This work introduces capillary electrophoresis &#8211; mass spectrometry (CESI-MS) as a new technique for NM corona characterization. The protocols outlined here demonstrate how CESI-MS can be used for the reliable characterization of both the protein and metabolite corona acquired by NMs. The move to CESI-MS greatly decreases the volume of sample required (compared to traditional liquid chromatography &#8211; mass spectrometry (LC-MS) approaches) with multiple injections possible from as little as 5 &#181;L of sample, making it ideal for volume limited samples. Furthermore, the environmental consequences of analysis are reduced with respect to LC-MS due to the low flow rates (&#60;20 nL/min) in CESI-MS, and the use of aqueous electrolytes which eliminates the need for organic solvents.

Here we present a protocol to characterize the complete biomolecular corona, proteins, and metabolites, acquired by nanomaterials from biofluids using a capillary electrophoresis &#8211; mass spectrometry approach.

The adsorption of biomolecules from surrounding biological matrices to the surface of nanomaterials (NMs) to form the corona has been of interest for the ...

Covalent Labeling with Diethylpyrocarbonate for Studying Protein Higher-Order Structure by Mass Spectrometry

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool for this purpose, especially for protein systems that are difficult to study by traditional methods. To study a protein&#39;s structure by MS, specific chemical reactions are performed in solution that encode a protein&#39;s structural information into its mass. One particularly effective approach is to use reagents that covalently modify solvent accessible amino acid side chains. These reactions lead to mass increases that can be localized with residue-level resolution when combined with proteolytic digestion and tandem mass spectrometry. Here, we describe the protocols associated with use of diethylpyrocarbonate (DEPC) as a covalent labeling reagent together with MS detection. DEPC is a highly electrophilic molecule capable of labeling up to 30% of the residues in the average protein, thereby providing excellent structural resolution. DEPC has been successfully used together with MS to obtain structural information for small single-domain proteins, such as &#946;2-microglobulin, to large multi-domain proteins, such as monoclonal antibodies.

The experimental procedures for performing diethylpyrocarbonate-based covalent labeling with mass spectrometric detection are described. Diethylpyrocarbonate is simply mixed with the protein or protein complex of interest, leading to the modification of solvent accessible amino acid residues. The modified residues can be identified after proteolytic digestion and liquid chromatography/mass spectrometry analysis.

Characterizing a protein&#39;s higher-order structure is essential for understanding its function. Mass spectrometry (MS) has emerged as a powerful tool ...