Accelerated Barocycler Lysis and Extraction ... - ACS Publications

Subscriber access provided by Stockholm University Library

Article

Accelerated Barocycler Lysis and Extraction (ABLE) sample preparation for clinical proteomics by mass spectrometry Natasha Lucas, Andrew B. Robinson, Maiken Marcker Espersen, Sadia Mahboob, Dylan Xavier, Jing Xue, Rosemary L. Balleine, Anna deFazio, Peter G Hains, and Phillip J Robinson J. Proteome Res., Just Accepted Manuscript • DOI: 10.1021/acs.jproteome.8b00684 • Publication Date (Web): 16 Nov 2018 Downloaded from http://pubs.acs.org on November 19, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Journal of Proteome Research

Accelerated Barocycler Lysis and Extraction (ABLE) sample preparation for clinical proteomics by mass spectrometry

Natasha Lucas1, Andrew B Robinson1, Maiken Marcker Espersen2, Sadia Mahboob1, Dylan Xavier1, Jing Xue3, Rosemary L. Balleine 1, Anna deFazio2,4,5, Peter G Hains1,3, Phillip J Robinson1,3

1.

ProCan®, Children’s Medical Research Institute, Faculty of Medicine and Health, University of Sydney, Westmead, NSW, Australia.

2.

Centre for Cancer Research, Westmead Institute for Medical Research, Westmead, NSW, Australia.

3.

Cell Signalling Unit, Children's Medical Research Institute, The University of Sydney

4.

Faculty of Medicine and Health, The University of Sydney, NSW, Australia.

5.

Department of Gynaecological Oncology, Westmead Hospital, Sydney, NSW, Australia.

ORCID: NL: 0000-0002-7656-6866 ABR: 0000-0001-7729-0331 MME: 0000-0003-1042-7342 SM: 0000-0002-1143-8935 DX: 0000-0003-2601-9343 RLB: 0000-0003-2864-4345 AdeF: 0000-0003-0057-4744 1 ACS Paragon Plus Environment

Journal of Proteome Research 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

PGH: 0000-0002-7276-1760 PJR: 0000-0002-7878-0313

Abstract We have developed a streamlined proteomic sample preparation protocol termed Accelerated Barocycler Lysis and Extraction (ABLE), that substantially reduces the time and cost of tissue sample processing. ABLE is based on pressure cycling technology (PCT) for rapid tissue solubilisation and reliable, controlled proteolytic digestion. Here, a previously reported PCT based protocol was optimised using 1-4 mg biopsy punches from rat kidney. The tissue denaturant urea was substituted with a combination of sodium deoxycholate (SDC) and N-propanol. ABLE produced comparable numbers of protein identifications in half the sample preparation time, being ready for MS injection in three hours compared with six hours for the conventional urea based method. To validate ABLE, it was applied to a diverse range of rat tissues (kidney, lung, muscle, brain, testis), human HEK 293 cell lines and human ovarian cancer samples, followed by SWATH-mass spectrometry (SWATH-MS). There were similar numbers of quantified proteins between ABLE-SWATH and the conventional method, with greater than 70% overlap for all sample types, except muscle (58%). The ABLE protocol offers a standardised, high-throughput, efficient and reproducible proteomic preparation method, that when coupled with SWATH-MS, has the potential to accelerate proteomics analysis to achieve a clinically relevant turn-around-time.

2 ACS Paragon Plus Environment

Page 2 of 23

Page 3 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Keywords Proteomics, Sample preparation, Barocycler, PCT, SWATH, Mass spectrometry, Tissue biopsy, Cancer, Clinical proteomics

Introduction Clinical application of tissue proteomics holds promise for substantial advances in the diagnosis and treatment of complex diseases. To achieve this, it is important that efficient workflows and standardised operating procedures are developed that can support delivery of reliable and accurate results in a clinically relevant timeframe using small amounts of sample1. Recent large-scale, systematic measurements of proteomes by sequential window acquisition of all theoretical fragment ion spectra - mass spectrometry (SWATH-MS) have generated large datasets that can be mined to determine the state of the proteome and its response to perturbations.

SWATH-MS has comparable reproducibility, linear dynamic range, and

sensitivity to selective reaction monitoring (SRM), without the need for a pre-determined set of peptides and transitions, allowing for relative quantitation across large sample sets2. Such data sets have revealed systemic malfunctions at the cellular and organismal levels in diverse diseases 3-4. Proteomics on a large scale requires sample preparation methods that are robust and reproducible as well as streamlined and suitable for high-throughput. In clinical proteomics, two main types of samples are analysed: body fluids (serum, CSF, tears, urine etc.) and solid tissues (tissue biopsies). There are up to two initial steps: tissue solubilisation for solid tissues, followed by proteolytic digestion of the released proteins. Solubilisation of solid tissues is the most labour-intensive step and one of the most variable steps that can impact on large scale studies. Traditional protocols for proteolytic digestion 3 ACS Paragon Plus Environment


involve overnight digestion with trypsin, with or without the second protease Lys-C. This long reaction time creates a workflow bottleneck, lacks consistent timing, and risks introduction of artificial protein modifications that can increase sample variability5. Options available to decrease the reaction time include microwave, ultrasound6, immobilised trypsin, infrared, solvent effects, heat, and filter-aided digestion. These afford little advantage over conventional overnight digestion7. However, the introduction of pressure cycling technology (PCT) using Barocycler instrumentation has greatly improved both tissue solubilisation and digestion consistency8-14. The Barocycler instrument cycles between high pressure (up to 45 kpsi) and ambient pressure approximately every minute to lyse tissues or cells, solubilise, extract and denature proteins, using a Dounce-like homogenisation principle, but based on pressure11. After solubilisation, a second round of pressure cycling is applied in the presence of trypsin/Lys-C for digestion, without the need to transfer samples to a new tube. Protein digestion is accelerated under high pressure9, 15-16. This allows for unambiguous control of digestion parameters, standardising digestion times, limits off-target reactions and provides a higher sequence coverage compared with atmospheric pressure10. As little as 0.2 mg (wet weight) of tissue can be processed, with samples ready for MS analysis in 6-12 hours13. PCT was first combined with SWATH-MS by Guo et. al. 201513.

PCT-SWATH-MS showed that complex proteome maps could be

reproducibly generated from a small 1-4 mg tissue biopsy in an overall 12-hour time frame13. The PCT-based sample preparation protocols reported to date generally utilise urea as a lysis buffer for protein denaturing and solubilisation11-13. Urea is a widely-used lysis buffer in shotgun proteomics, but has several drawbacks. Concentrations of >1 M urea are detrimental to trypsin activity17; it can introduce unwanted modifications like carbamylation of N-termini and lysine residues, especially with prolonged incubation, and when used at elevated temperatures18, urea is incompatible with a majority of standard protein quantification assays. 4 ACS Paragon Plus Environment

Page 4 of 23

Page 5 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Urea is used in high 6-8 M concentrations, which require desalting prior to MS injection. An alternative, sodium dodecylsulfate (SDS), is a well-known and powerful tissue and protein solubiliser, but is incompatible with LC-MS/MS analysis, can affect trypsin activity and is difficult to remove downstream. In contrast, the bile salt surfactant sodium deoxycholate (SDC) is an alternative detergent to SDS that can be easily removed with an acid precipitation step19-20. It can be used at relatively high concentrations without affecting trypsin activity21. The aim of this study was to determine whether replacing urea with SDC would produce similar tissue digestion profiles and improve the PCT method for high efficiency and thus facilitate greater sample throughput.

Materials/Methods Cells and Tissues Rat tissues were harvested, with approval from the Animal Care and Ethics Committee for the Children's Medical Research Institute, Sydney, Australia (project number C116). Frozen tissues samples were taken using a 3 mm biopsy punch (Kai Industries, Seki City, Japan) and placed into microtubes (Pressure Biosciences, South Easton, MA, USA). The wet weight ranged from 0.9 – 7.1 mg. HEK293T cells (from Dr Timothy Adams, CSIRO Manufacturing, Parkville, Victoria, Australia) were adapted to grow in suspension in Freestyle 293 Expression medium (Life Technologies Australia, Scorsby, VIC, Australia) supplemented with 200 mg/L G418 (Life Technologies) using a humidified shaker incubator (37°C, 5% CO2, 130 rpm). The adapted HEK293T culture was maintained in Erlenmeyer shaker flasks and scaled up in a 20 L WAVE bioreactor (GE Healthcare, Silverwater, NSW, Australia), seeded at an initial working volume 5 ACS Paragon Plus Environment


of 5 L at a concentration of 0.8 × 106 viable cells/mL. To reduce shear stress, Pluronic F68 (Life Technologies) was added to the culture at 0.2% (w/v) final concentration. At a viable cell density of 3 × 106 cells/mL the culture was scaled up to 20 L and maintained at 37°C with a rocking speed of 25 rpm and rocking angle of 9°. The cells were harvested at ~5 × 106 cells/mL by centrifugation (1,500 x g, 10 min, 4°C), snap-frozen on liquid nitrogen and stored at -80°C. Frozen cell pellets were scraped using a small spatula into a Barocycler microtube (Pressure Biosciences). The wet weight ranged from 1.2 – 5.4 mg. Cryopreserved tissue samples from ten individuals with high grade serous ovarian carcinoma were obtained from the Gynaecological Oncology Biobank (GynBiobank) at Westmead Hospital, Sydney. GynBiobank participants provide written informed consent for the use of bio-specimens in research. The study was approved by Western Sydney Local Health District Human Research Ethics Committee (reference: AU RED LNR/16/WMEAD/291). To ensure that samples submitted for comparative processing were as similar as possible, each tumour was initially cryo-sectioned and stained with haematoxylin and eosin (H&E) to confirm tissue content. Two morphologically similar areas were identified and 1.5 mm diameter cores were taken using the CryoXtract CXT350 (Bio-Strategy, Broadmeadows, VIC, Australia). Post-coring, a second H&E stained section was then taken to verify cored areas. Frozen tissue cores were divided and the surface segment submitted for processing. The wet weight of processed samples ranged from 0.6 – 2 mg. Tissue pieces were washed in microtubes to remove the optimal cutting temperature compound (OCT) prior to lysis as described by Zhang et. al.22, with the following modifications. Ice-cold 70% (v/v) ethanol in HPLC-grade water (120uL) was added to the tissue biopsy in a microtube and gently shaken for 1 min, followed by centrifugation for 1 min (5000 × g). The supernatant was carefully removed and the above step was repeated, followed by a final ice-cold HPLC-grade water wash, using the same conditions as above. 6 ACS Paragon Plus Environment

Page 6 of 23

Page 7 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Conventional PCT Method Tissue lysis, reduction and alkylation were carried out simultaneously using 30 µL of 6 M Urea, 50 mM ammonium bicarbonate, 10 mM tris(2-carboxyethyl)phosphine (TCEP) and 40 mM iodoacetamide (IOA) in a Barocycler 2320EXT (Pressure Biosciences) at 45 kpsi, for 60 cycles (50 s high pressure, 10 s atmospheric pressure). Following this, 5 µg of Lys-C endopeptidase (Novachem, Heidelberg West, VIC, Australia) was added to each sample tube and digestion was carried out in the Barocycler using 45 cycles of 50 s at 20 kpsi and 10 s at atmospheric pressure. After LysC digestion the urea concentration was diluted to 1.1 M with 50 mM ammonium bicarbonate and trypsin digestion was performed in the Barocycler for 90 cycles (50 s at 20 kpsi, 10 s at atmospheric pressure). All Barocycler steps were performed at 35⁰C. Following digestion, samples were desalted via C18 SPE columns (Empore SPE 4 mm/1 mL C18-SD) and the eluent dried. Samples were resuspended in 0.1% (v/v) formic acid and concentration was determined using A280 nm with a nanophotometer N60 (Implen, LabGear, Brisbane, QLD, Australia). One mg of wet weight typically resulted in 10-14 μg of peptide digest, based on nanophotometer analysis.

Optimised ABLE Method Tissue lysis, reduction and alkylation were carried out simultaneously using 30 µL of 1% (w/v) sodium deoxycholate (SDC), 5% (v/v) N-propanol, 100 mM triethylammonium bicarbonate, 10 mM tris(2-carboxyethyl)phosphine (TCEP) and 40 mM iodoacetamide (IOA in a Barocycler 2320EXT at 45 kpsi, for 60 cycles (50 s high pressure, 10 s atmospheric pressure), at 56⁰C. Following this, 120 µL of rapid digestion buffer (Promega) was added to each sample and 1 µg of rapid trypsin/Lys-C (Promega). Digestion was carried out in the 7 ACS Paragon Plus Environment


Barocycler using 30 cycles of 50 s at 45 kpsi and 10 s at atmospheric pressure, at 70⁰C. Samples were acidified with formic acid to precipitate the SDC, then centrifuged (15 min 18,000 x g). The supernatant was transferred to a new tube and evaporated to dryness. Samples were resuspended in 0.1% (v/v) formic acid and the concentration determined using A280 nm with a Implen nanophotometer N60. 1 mg of wet weight typically resulted in 10-14 μg of peptide digest.

LC-MS/MS Methods IDA Acquisition An Eksigent nanoLC 425 HPLC operating in microflow mode, coupled online to a 6600 Triple TOF (SCIEX) was used for the analyses. The peptide digests (2 µg) were spiked with retention time standards and injected onto a C18 trap column (SGE TRAPCOL C18 G203 300 µm x 100 mm) and desalted for 5 min at 10 µL/min with solvent A (0.1% [v/v] formic acid). The trap column was switched in-line with a reversed-phase capillary column (SGE C18 G203 250 mm × 300 µm ID 3 µm 200 Å), maintained at a temperature of 40⁰C. The flow rate was 5 µL/min. The gradient started at 2 % solvent B (99.9% [v/v] acetonitrile, 0.1% [v/v] formic acid) and increased to 10% over 5 min. This was followed by an increase of solvent B to 25% over 60 min, then a further increase to 40% for 5 min. The column was washed with a 4 min linear gradient to 95% solvent B held for 5 min, followed by a 9 min column equilibration step with 98% solvent A. The LC eluent was analysed using the Triple TOF 6600 system (SCIEX) equipped with a DuoSpray source and 50 µm internal diameter electrode and controlled by Analyst 1.7.1 software. The following parameters were used: 5500 V ion spray voltage; 25 nitrogen curtain gas; 100°C TEM, 20 source gas 1, 20 source gas 2. The 90 min information 8 ACS Paragon Plus Environment

Page 8 of 23

Page 9 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


dependent acquisition (IDA), consisted of a survey scan of 200 ms (TOF-MS) in the range 350–1250 m/z to collect the MS1 spectra and the top 40 precursor ions with charge states from +2 to +5 were selected for subsequent fragmentation with an accumulation time of 50 ms per MS/MS experiment for a total cycle time of 2.3 s and MS/MS spectra were acquired in the range 100–2000 m/z.

SWATH Acquisition For the SWATH acquisition, peptide spectra were acquired with the LC MS/MS method as described for IDA acquisition, using 100 variable windows, as per SCIEX technical notes (available at https://sciex.com/community/Asset/00001409/vw100_ces_5_10.txt).

The

parameters were set as follows: lower m/z limit 350; upper m/z limit 1250; window overlap (Da) 1.0; CES was set at 5 for the smaller windows, then 8 for larger windows; and 10 for the largest windows. MS2 spectra were collected in the range of m/z 100 to 2000 for 30 ms in high resolution mode and the resulting total cycle time was 3.2 s.

Spectral Reference Library Generation To decode SWATH spectra an independently generated spectral reference library (SRL) produced in IDA mode is required, ideally obtained from the same instrument and with the same HPLC gradient. To produce three SRLs pooled digests of rat tissues, HEK293T cells and ovarian cancer tissues were each pre-fractionated using the Pierce High pH Reversed-Phase Fractionation Kit (Thermo Fisher Scientific) following manufacturer’s instructions. The fractions were analysed by LC-MS/MS in IDA mode as described above. The rat tissue library was generated by combining samples of multiple tissue types, that were fractionated using high pH, and running ten fractions using 90 min IDA runs. The spectra were combined and searched 9 ACS Paragon Plus Environment


using Protein Pilot as described below. The resultant rat tissue library contained 4,457 proteins at 1% FDR. The HEK293T library and ovarian cancer library were produced in a similar manner and contained 4,692 and 5,404 proteins at 1% FDR, respectively. Protein pilot group files were converted to spectral library text files using PeakView software (SCIEX) v2.2. Retention time standards were selected within PeakView before the data was analysed on OneOmics (SCIEX).

Data Processing IDA spectra were searched using Protein Pilot v5.0 (SCIEX) using the Paragon algorithm with the following parameters: Sample Type: Identification; Cys Alkylation: Iodoacetamide; Digestion: Trypsin; Instrument: TripleTOF 6600; Database: Uniprot Rat (37,596 entries) for all rat tissues, Uniprot Human (178,750 entries) for HEK293T and ovarian cancer samples. Both databases contained sequences for internal retention time calibration standards. Thorough ID and False Discovery Rate (FDR) Analysis were selected, and the FDR was set at 1%. The SWATH files and ion library files were uploaded to BaseSpace (Illumina, San Diego, CA, USA) via CloudConnect Software v1.0 (SCIEX) on PeakView v2.2 (SCIEX). Samples were processed using the OneOmics package, starting with the Protein Expression Extractor application v1.0.0 (SCIEX), with the following parameters: Number of peptides per protein: 6; Number of transitions per peptide: 6; XIC extraction window (in minutes): 5; XIC extraction widths: 40; XIC extraction widths unit: ppm. Results were further processed using the Protein Expression Workflow application v1.1.0 Beta (SCIEX), and exported via the Analytics tab. Calculation of peptide physicochemical properties was performed with R using the 'Peptides' package23. See Supplementary Methods for coding details. 10 ACS Paragon Plus Environment

Page 10 of 23

Page 11 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The data (significantly impacted pathways, biological processes, molecular interactions, miRNAs, SNPs, etc.) were analysed using iPathwayGuide, Advaita Bioinformatics (http://www.advaitabio.com/ipathwayguide). The fold change output from OneOmics was uploaded to iPathway Guide and differentially expressed protein thresholds with a log foldchange of at least 0.6 and p-value ≤ 0.05 were used. Elim pruning was used to correct pvalues for gene ontology (GO) terms and false discovery rate (FDR) was used for pathway analysis correction.

Results and Discussion To evaluate the suitability of SDC for potentially accelerating sample preparation, a series of experimental conditions was assessed using rat kidney to determine the final optimised sample preparation parameters. Variables included; the concentration of SDC, addition of Npropanol to lysis buffer, one-step and two-step digestions, lysis temperature and time. The final optimised protocol was termed Accelerated Barocycler Lysis & Extraction (ABLE) (Figure 1).

Figure 1. Method flow diagram for conventional PCT-SWATH workflow (a) compared to the optimised Accelerated Barocycler Lysis & Extraction (ABLE) ABLE-SWATH method (b).



ABLE comprised two key steps in the Barocycler, firstly for tissue lysis and secondly for proteolytic digestion, compared with three steps for the conventional urea-based method. The lysis step, was carried out using a combination of 1% (w/v) SDC and 5% (v/v) N-propanol. The addition of N-propanol improved digestion efficiency and reproducibility when using the regular two-step digestion method (LysC for 60 cycles, followed by trypsin 90 cycles), however upon employing Promega Rapid Trypsin/LysC, N-propanol added little specific value (Supplementary Figure S1). The switch to SDC resulted in essentially no carbamylation of the N-terminus or Lys residues (Supplementary Figure 2). SDC allowed the use of higher temperatures compared to urea, and hence the first step (lysis, reduction and alkylation) was performed at 56℃. Although 30 cycles for lysis was sufficient for kidney samples (Supplementary Figure S3a), in the final protocol this lysis step was set


Page 12 of 23

Page 13 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


at 60 cycles to allow for lysis of solid tumour samples which can be quite variable. Unpublished work has shown difficult samples, including muscle, are well homogenised with 30, 45 or 60 min lysis with our methodology (data not shown). Next, the second digestion step in the Barocycler was optimised and the third step was eliminated. The conventional urea-based method involves digestion in two stages, with Lys-C for 45 cycles, followed by dilution of the urea and trypsin digestion for a further 60 cycles (because urea inhibits trypsin unless the urea is sufficiently diluted). This additional dilution step increases analysis time and the chance of manual error. Since trypsin tolerates SDC, the dilution step is not required with ABLE. To further reduce digestion time, we capitalised on Rapid Trypsin/Lys-C from Promega. This maintained robust activity at 70⁰C, and allowed for Barocycler digestion to be performed in a single step, completing digestion in 30 cycles (approx. 30 min) rather than 105 min (Supplementary Figure S3b). We do see more artefactual modification of some amino acids with the increased temperature (70⁰C vs 37⁰C, Supplementary Figure 4a), however this is largely in-line with standard overnight digestion protocols24. Furthermore, the level of modified peptide is generally a low percentage of the total (native plus modified) peptide in the sample. The change from 20 kpsi to 45 kpsi for the 1-step with Rapid Trypsin/Lys-C (Supplementary Figure 4b) improves digestion compared to the 2-step procedure at 20 kpsi. There is an arguable improvement in digestion for Rapid Trypsin/Lys-C at 20 kpsi versus 45 kpsi. SDC can easily be removed from the final digested sample with an acid precipitation step. As the sample contains low salt solutions, desalting can be done on line in the LC-MS using a trap-elute set-up, avoiding the need and associated cost for a separate off-line C18 solid phase extraction (SPE) clean-up. An SPE clean-up was not used for any of the samples prepared for this report, but it remains unclear as yet if this can be avoided for all tissue types. 13 ACS Paragon Plus Environment


To validate the final ABLE protocol, both ABLE-SWATH and urea-based PCT-SWATH was applied to a diverse range of rat tissues: kidney, testis, brain, lung and muscle. To demonstrate the versatility of the method, human HEK293T cell lines and human ovarian cancer samples were also prepared. The rat and HEK293 samples were prepared in two batches of four biological replicates for each method and each batch was run on separate MS instruments using identical conditions. Ten ovarian cancers were sampled in duplicate with a single sample submitted for each method, and two technical replicates of each extract run on one single MS instrument. Since all tumour samples were biological replicates and each tissue biopsy (~2 mg) was punched and placed in separate tubes, this leads to larger data variation, particularly for the tumour samples relative to the rat organs. Each sample was searched against the relevant SRL. For all tissues except muscle, ~4,000 peptides were quantified by SWATH-MS on average per sample, meeting the 1% FDR cut-off (Figure 2). Overall, the optimised method showed highly comparable numbers of quantified proteins and peptides with a similar reproducibility for each biological replicate, regardless of the sample tissue source.

Figure 2. Comparison of the performance of ABLE-SWATH with conventional PCTSWATH. Average number of quantified peptides per sample in various rat tissues (n=8 ± SEM), HEK293T cells (n=8 ± SEM) and human ovarian tumour samples (n=20 ± SEM) at 1% FDR.


Page 14 of 23

Page 15 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The conventional and ABLE methods each gave an average of >70% identity overlap for quantified proteins (Figure 3). Approximately 50% peptide overlap (summed biological replicates, rat tissues (n=8), HEK293T cells (n=8) and ovarian tumour samples (n=10)) was seen at 1% FDR (Supplementary Figure S5).

Figure 3. Overlap between proteins identified with ABLE-SWATH and conventional PCTSWATH. Venn diagrams show the overlap of quantified proteins in any of the various rat tissues (n=8), HEK293T cells (n=8) and human ovarian tumour samples (n=10) filtered at 1% FDR.



The overlap of all the rat tissues identified by the ABLE-SWATH method are shown in Supplementary Figure S6 and the unique proteins found in each tissue type shown in Supplementary Figure S6 are listed in Supplementary Table 1. This demonstrates the ability of SWATH-MS to discriminate tissue types, with 1,506 proteins being fully in common to all five rat organ types, while 41-145 proteins were wholly unique at this sensitivity level to a single organ. The proteolytic digestion efficiency between the two methods was assessed using IDA runs generated for each sample type and was found to be comparable (Figure 4), indicating no digestion bias in the ABLE method. All tissues (except muscle) showed around 70% canonical tryptic sequences. Muscle produced and identified fewer peptides for either method and resulted in 60% fully tryptic peptides, potentially due to the impact of high abundance proteins of the contractile machinery (e.g. myosin, actin and nebulin).

Figure 4. Digestion efficiency for each sample type, using the optimised ABLE method or the conventional PCT preparation for rat tissues (n=8) and HEK293T cells (n=8).


Page 16 of 23

Page 17 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Since ABLE-SWATH revealed 9-19% unique proteins relative to conventional PCTSWATH we asked whether there were systematic differences in the ABLE-SWATH method or whether this may represent expected biological variability. We investigated whether any extraction differences might exist between the two methods based on physicochemical properties of the peptides. Calculations on the peptide hydrophobicity, isoelectric point (pI) and charge were performed on the peptides found to be unique to either the ABLE-SWATH or the conventional method, to investigate if there was a physicochemical influence on each methods’ peptide selectivity. No significant physicochemical property differences were found between the unique peptides in the two methods across all the rat tissues (Supplementary Figure S7). We determined whether there was a bias to any particular cellular pathway between the two methods. Pathway impact analysis was performed on the proteins identified in the five different rat tissues. The analysis was done using those proteins found commonly across both 17 ACS Paragon Plus Environment


SWATH methods, but which were significantly increased or decreased in abundance in one or other method (p < 0.05, log 10 fold change 0.6). For the tissues kidney, brain, testis, muscle and lung, 439, 595, 709, 316, and 521 differentially expressed proteins were identified respectively, indicating that the two methods show differences even within the proteins detected by both. Of these proteins, only 19 displayed consistent differential abundance across all tissues (Supplementary Figure S8), six of which were more abundant in the ABLE method and 13 more abundant in conventional PCT-SWATH. The pathway analysis showed no significantly affected pathways across all tissues. Gene ontology analysis, for biological processes, molecular functions and cellular components, found only one significant GO term (molecular function) ‘structural constituents of ribosome’. This suggests that the differences between relative quantitation in the methods are not systematically biased to recovery and identification of different proteins in different GO pathways, but more likely reflect a combination of biological and instrument variability. To demonstrate the utility of the method for clinical specimens, ten human ovarian cancer samples were prepared by taking two comparable cores from a single specimen (Supplementary Figure S9a). The conventional and optimised ABLE methods identified similar numbers of peptides at 1% FDR for the majority of cancer samples. There was a greater degree of variation between paired samples compared with rat tissues, which may be attributable to greater tissue heterogeneity (Supplementary Figure S9b). Overlap between proteins identified in the ovarian cancer samples prepared by the two methods was 83.5%, the highest of any of the sample types measured (Figure 3). These results demonstrate that the optimised ABLE method based on SDC was comparable to the conventional PCT method based on urea in digestion efficiency, protein and peptide identification and reproducibility. The use of SDC in the lysis buffer gave greater flexibility for optimisation of the existing PCT workflow, allowing higher temperatures to be used, and 18 ACS Paragon Plus Environment

Page 18 of 23

Page 19 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


in turn, reducing the number of Barocycler cycles required, particularly for the enzymatic digestion step. The method minimised the number of sampling handling steps, which will ultimately reduce user error and technical variation, whilst improving sample throughput. Omitting the need for offline sample clean-up will also prevent further issues with variation and recovery and save time, although this needs validation across a larger and more diverse tissue set. The cost of the method per sample is also decreased, due to reduced consumable requirements (extra PCT caps and SPE cartridges). This can become considerable, when processing samples on a large scale. The greatest advantage of the optimised method is that samples are processed by ABLE in only 2-3 hours versus 6-8 hours for the conventional urea method.

Conclusion Our data present an improved conventional tissue PCT approach in a Barocycler by replacing urea and proteolytic enzymes with SDC, N-propanol, and modified commercially available enzymes that have higher optimum temperatures. The ABLE method is achieved in half the time of the original method, using fewer steps, hence improving throughput and reducing the potential for error. The final method uses a two-step Barocycler approach of 60 cycles for lysis and 30 cycles for digestion. This resulted in a method which produces comparable results to the conventional three-step PCT method based on urea. It is a reliable, broad utility, sample preparation method that can be completed within a clinically relevant time frame and which also allows for high-throughput clinical cohort studies at reduced costs without compromising quality.



Acknowledgements The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE25 partner repository with the dataset identifier PXD010688. Authors wish to thank Dr. Vera S. Gross for consulting on the Barocycler optimisation and data on improved trypsin and Lys-C efficiency under high pressure and Dr. Scott Cohen for providing the HEK293 cell line samples. This work was supported by equipment grants from the Australian Cancer Research Foundation (ACRF) and project grants from the Office for Health and Medical Research (OHMR) NSW, Australia, the Cancer Institute NSW (CINSW) as a Translational Program Grant (TPG), and from the National Health & Medical Research Council (NHMRC) Australia (GNT1069493, GNT1047070, and GNT1032771). M.M.E. is recipient of a Cancer Institute NSW Early Career Fellowship. The Gynaecological Oncology Biobank at Westmead is a member of the Australasian Biospecimen Network-Oncology group and has been supported by the National Health and Medical Research Council of Australia, Enabling Grants ID 310670 and ID 628903 and the Cancer Institute NSW grants 12/RIG/1-17 and 15/RIG/1-16.


Page 20 of 23

Page 21 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


References 1.

Zhou, L.; Wang, K.; Nice, E.; Zhang, H.; Huang, C., Clinical proteomics-driven precision medicine

for targeted cancer therapy: current overview and future perspectives. Expert Review of Proteomics 2016, 13 (4), 367-381. 2.

Collins, B. C.; Hunter, C. L.; Lui, Y.; Schilling, B.; Rosenberger, G.; Bader, S. L.; Chan, D. W.;

Gibson, B. W.; Gingras, A.; Held, J. M.; Hirayama-Kurogi, M.; Hou, G.; Krisp, C.; Larsen, B.; Lin, L.; Liu, S.; Molloy, M. P.; Moritz, R. L.; Ohtsuki, S.; Schlapbach, R.; Selevsek, N.; Thomas, S. N.; S. Tzeng; Zhang, H.; Aebersold, R., Multi-laboratory assessment of reproducibility, qualitative and quantitative performance of SWATH-mass spectrometry. Nature Communications 2017, 8 (1), 291. 3.

Aebersold, R.; Mann, M., Mass-spectrometric exploration of proteome structure and function.

Nature 2016, 537 (7620), 347-55. 4.

Aebersold, R.; Mann, M., Mass spectrometry-based proteomics. Nature 2003, 422 (6928),

198-207. 5.

Klammer, A. A.; MacCoss, M. J., Effects of Modified Digestion Schemes on the Identification

of Proteins from Complex Mixtures. Journal of Proteome Research 2006, 5 (3), 695-700. 6.

Lo´pez-Ferrer, D.; Capelo, J. L.; Va´zquez, J., Ultra Fast Trypsin Digestion of Proteins by High

Intensity Focused. Journal of Proteome Research 2005, 4 (5), 1569-1574. 7.

Hustoft, H. K.; Reubsaet, L.; Greibrokk, T.; Lundanes, E.; Malerod, H., Critical assessment of

accelerating trypsination methods. Journal of Pharmaceutical and Biomedical Analysis 2011, 56 (5), 1069-1078. 8.

Gross, V.; Carlson, G.; Kwan, A. T.; Smejkal, G.; Freeman, E.; Ivanov, A. R.; Lazarev, A., Tissue

fractionation by hydrostatic pressure cycling technology:The unified sample preparation technique for systems biology studies. Journal Biomolecular Techniques 2008, 19 (3), 189-199.



9.

Freeman, E.; Ivanov, A. R., Proteomics under Pressure: Development of Essential Sample

Preparation Techniques in Proteomics Using Ultrahigh Hydrostatic Pressure. Journal of Proteome Research 2011, 10 (12), 5536-5546. 10.

Olszowy, P. P.; Burns, A.; Ciborowski, P. S., Pressure-assisted sample preparation for

proteomic analysis. Analytical Biochemistry 2013, 438 (1), 67-72. 11.

Shao, S.; Guo, T.; Gross, V.; Lazarev, A.; Koh, C. C.; Gillessen, S.; Joerger, M.; Jochum, W.;

Aebersold, R., Reproducible Tissue Homogenization and Protein Extraction for Quantitative Proteomics Using MicroPestle-Assisted Pressure-Cycling Technology. Journal of Proteome Research 2016, 15 (6), 1821-1829. 12.

Shao, S.; Guo, T.; Koh, C. C.; Gillessen, S.; Joerger, M.; Jochum, W.; Aebersold, R., Minimal

sample requirement for highly multiplexed protein quantification in cell lines and tissues by PCTSWATH mass spectrometry. Proteomics 2015, 15 (21), 371-3721. 13.

Guo, T.; Kouvonen, P.; Koh, C. C.; Gillet, L. C.; Wolski, W. E.; Röst, H. L.; Rosenberger, G.; Collins,

B. C.; Blum, L. C.; Gillessen, S.; Joerger, M.; Jochum, W.; Aebersold, R., Rapid mass spectrometric conversion of tissue biopsy samples into permanent quantitative digital proteome maps. Nature Medicine 2015, 21 (4), 407-413. 14.

Powell, B. S.; Lazarev, A. V.; Carlson, G.; Ivanov, A. R.; Rozak, D. A., Chapter 2: Pressure Cycling

Technology in Systems Biology; Microbial Systems Biology: Methods and Protocols. In Methods in Molecular Biology, 2012; Vol. 881, pp 27-62. 15.

Lo´pez-Ferrer, D.; Petritis, K.; Hixson, K. K.; Heibeck, T. H.; Moore, R. J.; Belov, M. E.; Camp, D.

G.; Smith, R. D., Application of Pressurized Solvents for Ultrafast Trypsin Hydrolysis in Proteomics: Proteomics on the Fly. Journal of Proteome Research 2008, 7 (8), 3276-3281. 16.

Alvarado, R.; Tran, D.; Ching, B.; Phinney, B. S., A Comparative Study of In-Gel Digestions Using

Microwave and Pressure-Accelerated Technologies. Journal of Biomolecular Techniques 2010, 21 (3), 148-155.


Page 22 of 23

Page 23 of 23 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


17.

Beynon, R. J.; Bond, J. S., Appendix II “Commercially Available Proteases”. In Proteolytic

Enzymes, A Practical Approach, IRL Press: Oxford, U.K., 1989. 18.

Kollipara, L.; Zahedi, R. P., Protein carbamylation: In vivo modification or in vitro artefact?

Proteomics 2013, 13 (6), 941-944. 19.

Proc, J. L.; Kuzyk, M. A.; Hardie, D. B.; Yang, J.; Smith, D. S.; Jackson, A. M.; Parker, C. E.;

Borchers, C. H., A Quantitative Study of the Effects of Chaotropic Agents, Surfactants, and Solvents on the Digestion Efficiency of Human Plasma Proteins by Trypsin. Journal of Proteome Research 2010, 9 (10), 5422-5437. 20.

León, I. R.; Schwämmle, V.; Jensen, O. N.; Sprenger, R. R., Quantitative assessment of in-

solution digestion efficiency identifies optimal protocols for unbiased protein analysis. Molecular Cellular Proteomics 2013, 12 (10), 2992-3005. 21.

Lin, Y.; Zhou, J.; Bi, D.; Chen, P.; Wang, X.; Liang, S., Sodium-deoxycholate-assisted tryptic

digestion and identification of proteolytically resistant proteins. Analytical Biochemistry 2008, 377 (2), 259-266. 22.

Zhang, W.; Sakashita, S.; Taylor, P.; Tsao, M. S.; Moran, M. F., Comprehensive proteome

analysis of fresh frozen and optimal cutting temperature (OCT) embedded primary non-small cell lung carcinoma by LC–MS/MS. Methods 2015, 81, 50-55. 23.

Osorio, D.; Rondon-Villarreal, P.; Torres, R., Peptides: A package for data mining of

antimicrobial peptides. . The R Journal 2015, 7 (1), 4-14. 24.

Hains, P. G.; Robinson, P. J., The Impact of Commonly Used Alkylating Agents on Artifactual

Peptide Modification. Journal of Proteome Research 2017, 16 (9), 3443-3447. 25.

JA, V.; A, C.; N, d.-T.; JA, D.; J, G.; I, L.; G, M.; Y, P.-R.; F, R.; T, T.; QW, X.; R, W.; H, H., 2016

update of the PRIDE database and related tools. Nucleic Acids Res 2016, 44 (D1), D447-D456.


Accelerated Barocycler Lysis and Extraction ... - ACS Publications

Recommend Documents