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A Parallel Proteomic Workflow for Mass Spectrometric Analysis of Tissue Samples Preserved by Different Methods Ales Holfeld, Alberto Valdés, Per-Uno Malmström, Ulrika Segersten, and Sara Bergstrom Lind Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b00379 • Publication Date (Web): 06 Apr 2018 Downloaded from http://pubs.acs.org on April 9, 2018

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Analytical Chemistry

A Parallel Proteomic Workflow for Mass Spectrometric Analysis of Tissue Samples Preserved by Different Methods Aleš Holfeld1, Alberto Valdés1, Per-Uno Malmström2, Ulrika Segersten2, Sara Bergström Lind1* 1

Department of Chemistry-BMC, Analytical Chemistry, Uppsala University, Uppsala, Sweden. 2

Department of Surgical Sciences, Uppsala University, Uppsala, Sweden

*Corresponding author: Sara Bergström Lind, Department of Chemistry-BMC, Analytical Chemistry, Uppsala University, Box 599, 751 24 Uppsala, e-mail: [email protected], ORCID: 0000-0002-9510-3816

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Analytical Chemistry 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

Abstract Formalin-fixed and paraffin-embedded (FFPE) and optimal cutting temperature (OCT)-embedded and frozen tissue specimens in biobanks are highly valuable in clinical studies but proteomic and posttranslational modification (PTM) studies using mass spectrometry (MS) have been limited due to structural arrangement of proteins and contaminations from embedding material. This study aims to develop a parallel proteomic workflow for FFPE and OCT/frozen samples that allows for large-scale, quick, reproducible, qualitative and quantitative high-resolution MS-analysis. The optimized protocol gives details on removal of embedding material, protein extraction, and multi-enzyme digestion using filter-aided sample preparation method. The method was evaluated by investigating the protein expression levels in non-muscle-invasive and muscle-invasive bladder cancer samples in two cohorts and MS spectra were carefully reviewed for contaminations. More than 2000 and 3000 proteins in FFPE and OCT/frozen samples, respectively, were identified, and samples could be clustered in different tumour stages based on their protein expression. Furthermore, more than 250 and 400 phosphopeptides could be identified from specific patient samples of FFPE and OCT/frozen, respectively, using titanium dioxide enrichment. The paper presents unique data describing the similarities and differences observed in FFPE and OCT/frozen samples and shows the feasibility to detect proteins and site-specific phosphorylation even after long-term storage of clinical samples.

Keywords: Formalin-fixed and paraffin-embedded, Optimal cutting temperature embedded, Mass spectrometry, Proteomics, post-translational modifications


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Introduction Formalin-fixed and paraffin embedded (FFPE) and optimal cutting temperature (OCT)-embedded and cryopreserved tissue samples constitute a large part of repositories worldwide. The tradition of biobanking is of extreme value for clinical studies. Up to date, immunohistochemistry and other staining methods have been used in the biomedical analysis of such samples. Nowadays we have the possibility to perform global protein and post-translational modification (PTM) analysis of cells and tissue using mass spectrometric (MS) detection. For example, the widely used shotgun/bottom-up proteomics approach entails enzymatic digestion of proteins into peptides that are subsequently separated by liquid chromatography (LC) and analysed by MS and tandem MS/MS 1. Thereafter, MS/MS spectra of the surrogate peptides are used for database driven protein and post-translational modification identification. Most PTMs are generally low in abundance and their detection requires various enrichment strategies prior to LC-MS/MS analysis. For example, a number of different methods are available for the enrichment of phosphorylated peptides

2,3

. Currently, titanium dioxide (or analogue metal oxides) enrichments are

straightforward and efficient techniques due to the high affinity between the phosphate group and the material 3. In the past, there has been a notable barrier for investigating the samples in biobanks with LCMS/MS due to i) formalin has deleterious effects on protein structure and ii) embedding matrices, such as paraffin and OCT, which is a mixture of polymers (e.g., polyvinyl alcohol (PVA) and polyethylene glycol (PEG)), interfere with MS analysis if not properly removed. Therefore, the standard protocols for MSbased proteomics need to be further revised to enable analysis of these types of samples. Investigations of FFPE tissue samples recently gained attention, and tissues of diverse origin, such as breast, liver, kidney, colon, pancreas, prostate and urinary bladder have been explored by MS

4-10

. Current advances in sample

preparation, including heat-induced protein extraction and filter-aided sample preparation (FASP) method, have been milestones for the processing of FFPE samples

11

. Both quantitative proteomics and PTM

analysis in preserved tissues have been proven. For OCT samples, the concerns about ion suppression and



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contamination from polymers have been problematic, and proteomic studies by MS have, to the authors’ knowledge, only been reported in nine studies 7,12-19.

Many countries have a long tradition of storing large cohorts of samples in biobanks with related clinical data, which stresses the significance of novel developments in the sample preparation processes prior to MS experiments. There are some reports where both FFPE and OCT samples have been considered

13,14

,

but no study has reported such simultaneous investigation and comparison of FFPE and OCT samples as in this study. Here, we present the first optimized protocol for parallel analysis of FFPE and OCT embedded tissue samples using MS-based shotgun proteomics. Different protocols for i) embedding material removal, ii) protein extraction, and iii) multi-enzymatic digestion are developed and compared. The model system comprises a unique set of urothelial bladder cancer tissues that represented different stages of bladder cancer where samples from each patient have been preserved using both methods in parallel. A label-free approach for relative quantitation of protein expression profiles of two stages of cancer (non-muscle-invasive, T1, and muscle-invasive, T3), observed from the different sample types, is reported. In this way the preservation methods are also compared. Furthermore, maximized phosphopeptide enrichment from tissue sections from a single patient is carried out using titanium dioxide (TiO2) particles-based enrichment.

Experimental section Chemicals All chemicals were from Sigma-Aldrich (St Louis, MO, USA) if not otherwise stated. Materials The T24 cell line derived from highly malignant urinary bladder transitional-cell cancer was grown in RPMI-1640 cell medium (Life Technology, Grand Island, NA, USA) supplemented with 10% fetal bovine


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serum and 1% penicillin-streptomycin (both from Life Technology). Cells were grown to 80% confluence in a 150 cm2 flask before harvesting. Upon harvest, cell medium was removed and cells were washed with Dulcecco’s DPBS (Life Technology), detached with 0.25% trypsin incubation for 10 min and then physiological buffer saline (PBS) was added to collect the cells. Cells were washed with PBS three times and with DPBS two times. Finally, liquid was removed and cell pellet was stored in -80 °C. The PBS and DPBS solutions were supplemented with 1 mM sodium ortovanadate that served as a tyrosine phosphate inhibitor. A set of urinary bladder cancer tissue samples were provided from patients undergoing surgical resection between 2005 and 2009 at Uppsala University Hospital (Uppsala, Sweden). Ethical permission for using the material for MS analysis is granted by Uppsala University DNR 2015-143-1. Specimens collected were processed for preservation within 1-2 h after surgery. Parts from all specimens used were preserved with both OCT/freezing and FFPE procedures and from each approach we retrieved duplicate samples. For the OCT/frozen approach, samples were fixed in Tissue-Tek® O.C.T.TM compound (Sakura, The Netherlands) and snap frozen in liquid nitrogen. The OCT/frozen blocks were sectioned using a cryostat at temperature 75% were accepted.

Statistical analysis For statistical analysis, the “proteingroups.txt” files obtained from MaxQuant were loaded to the Perseus software (version 1.5.8.5). Potential contaminants, reverse hits, and proteins only identified by site were removed from the protein list, and the LFQ intensities were transformed into their log2-values. To assess the sample preparation reproducibility, log2 LFQ intensities of two biological replicates (from the same sample) were plotted against each other, and the correlation was evaluated according to the Pearson’s correlation coefficient. This correlation coefficient was also used to study the agreement between the different preservation methods. In this case, the two biological replicates were searched as one experiment and only proteins with all valid values of LFQ intensities in both OCT/frozen and FFPE tissues were considered. In addition, principal component analysis (PCA) was carried out to visualize the projection of the data sets. To determine differentially expressed proteins (DEP) between two distinct groups of patients 11 ACS Paragon Plus Environment


a two-tailed Student’s t-test was applied, considering significantly expressed those proteins with p < 0.05. Data were visualized using Volcano plots generated by plotting the log2 fold changes against the negative logarithm of the p-values. Safety Considerations Chemicals for sample preparation might be toxic and/or carcinogenic and biological material should be handled with great care according to the local laboratory guidelines.


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Results and discussion Development of protocols for MS-analysis of peptides and proteins in biobanked tissue is challenging but important in order to increase the use of clinical sample collections. In this study, we have developed a protocol that, to as considerable extent as possible, is overlapping for samples preserved by FFPE and OCT followed by cryopreservation. In this unique study, it was possible to study the impact of different preservation methods on protein expression in tumour tissues. It should be noted that samples had been stored in a biobank for >10 years and developments reported on here have been made to provide access to such long-term stored material using a simple protocol that can be used in clinic. The scheme of the present study is shown in Figure 1.

Optimization of sample preparation protocols. For OCT/frozen samples, most optimization concerned the removal of the fixation matrix. The main components, PVA and PEG, may not only cause ion suppression of detected peptides in the MS spectrum, but also substantially contaminate the LC-MS system. The establishment of a proper protocol was therefore tested out using MALDI experiments before samples were analysed by LC-MS/MS. To remove the OCT polymers, a procedure using ice-cold 70% ethanol, water, and washing buffer as proposed by other studies 17 was used. In Figure 2A, a total ion chromatogram (TIC) of an LC-MS/MS analysis of an initial attempt is presented. Even though the MALDI spectrum of this sample did not show any contaminants present (data not shown), the TIC profile revealed a series of intense polymer signals in the LC-MS/MS analysis. Furthermore, the chromatographic profile after in-solution digestion displayed a shift against hydrophobic/longer peptides, and the hydrophilic peptides were missing. When searching the data to identify the proteins, a high percentage (37%) of missed cleavages was observed, indicating insufficient digestion caused by the presence of polymers inhibiting the tryptic digestion. Therefore, the protocol was further developed, and it contains additional washing steps with ice-cold ethanol, water, and 13 ACS Paragon Plus Environment


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washing buffer as described in the Experimental section. By switching from in-solution to FASP digestion as proposed by Weston et al.16, it was also possible to include critical washing steps of the proteins prior to the addition of trypsin. Using the optimized procedure, the percentage of peptides with miscleavages was reduced from 37 to 13%. A TIC from the successful protocol is represented in Figure 2B, which is very similar to the one obtained from the reference sample (T24 cancer cell line) without embedding matrix (Figure 2E). In the case of FFPE samples, the protective paraffin wax had to be removed before proteins were extracted. The tissue pieces in current study were very small and surrounded by large excess of wax. This made it difficult to manually remove wax and instead, the removal relied on dissolvation using proper solvents since we were aiming for a protocol without the need for tools as laser-capture microdissection. This technology has been tested for FFPE samples as reported by e.g. Hood et al 7 with subsequent MSbased proteomic approach and proven to produce enough sample amounts for quantitative proteomics. Since microdissection is time consuming and not available to all clinical labs, the presented protocol in this paper is more straight-forward. In our initial trials, the utilization of heptane (proposed by Paulo et al. 23

) or xylene in combination with ethanol for rehydration and rinsing as in previous reports

8,9

, did not

provided satisfactory spectra in LC-MS/MS analysis since patterns of wax were observed in the chromatograms (Figure 2C). 1-D SDS-PAGE can be used to remove excess paraffin 4, but would require in-gel digestion, which have less peptide recovery. Instead, to overcome these difficulties, double solvents and a novel approach that uses the application of the so-called “hot-and-cold” cycle (proposed by Lai et al. 5

) were applied. In our optimized protocol xylene and heptane were used as primary and secondary

solvents, respectively and after applying heptane, an aliquot of methanol was added to create a two-phase system. Thereafter, the temperature was varied as described in the Experimental section. The chromatographic LC-MS/MS profile of a sample processed with this optimized protocol (Figure 2D) also overlapped with the profile observed from our cell line reference sample (Figure 2E), indicating that the procedure was adequate.


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For the protein extraction optimization, four different buffers were investigated (as specified in the Experimental section). Since the method development was performed on clinical samples of limited amounts, each buffer had to be tested on a new specimen. It was therefore not possible to evaluate all combinations of protein extraction and removal of embedding material procedures but on average 0.3 mg protein per mg tissue was extracted. The initial hypothesis was that detergent-containing buffers would be most efficient in extracting a wide range of proteins, e.g. also membrane proteins. However, such buffers required protocols that included removal steps for the detergents and it has been reported for OCT samples that precipitation with trichloroacetic acid or extraction with organic solvents are useful when detergent based buffers have been used for protein extractions18,19. In our case, samples evaluated with buffers C and D, i.e. containing OBG or SDS, were not free from detergents even though the FASP protocol or precipitation of the proteins were tested. A likely explanation is that when samples contained both contaminating polymeric embedding matrix and detergent in protein extraction buffer, the washing steps in the FASP and/or precipitation procedures were not enough to remove all species of polymeric origin. Instead, urea protein extraction buffers were evaluated (buffers A and B). Buffer B (based on urea, thiourea and a high concentration of ABC, 1M) clearly provided the highest yield for the OCT and FFPE samples tested. For FFPE samples a step with elevated temperature was used in the protein extraction to reverse the effect of formalin fixation, while for OCT samples the protein extraction was performed at low temperature. It has been previously observed that the use of urea and high temperature can generate carbamyl modifications on the proteins in FFPE samples due to the reaction of isocyanic acid with the side chains of lysine and arginine residues (isocyanic acid is derived from spontaneous dissociation of urea in an aqueous environment) 24. The addition of ABC has been suggested to prevent this reaction 24. Although we added this compound to our buffer, carbamylation was the most frequently observed protein modification, which could impede the digestion process of FFPE samples

24,25

. The effect of

carbamylation is further discussed below. Using these buffers for protein extraction no contaminations could be observed in the chromatograms using the T24 sample as reference (Figure 2B, 2D and 2E).



To improve the protein digestion efficacy, the optimized digestion approach was based on the consecutive digestion strategy (comprising two proteolytic enzymes Lys-C and trypsin), as suggested by Wisniewski et al.

10

. The most crucial optimization of the digestion procedure was to utilize the on-filter digestion

approach (FASP), since it enabled the more washing steps to remove contaminating preservatives that otherwise influenced the chromatographic pattern (Figure 2A and 2C).

Database search strategies Different database search strategies were used. Firstly, two biological replicates of the same tumour specimen from each preservation method were analysed and individually processed including or excluding carbamylation as variable modification. The number of identified proteins in OCT/frozen samples slightly increased (less than 3%) when this modification was included in the search (Table S1). However, this modification had a substantial impact on the total number of identified proteins in the FFPE samples, increasing the numbers by approximately 20% in most of the cases (Table S1). Secondly, when studying the agreement between the different preservation methods, the two biological replicates from the same preservation method were searched, including or excluding the carbamylation modification, as one experiment to increase the number of identifications (Table S2). Now the number of identified proteins slightly decreased in the OCT samples, slightly increased in the reference material and clearly increased in the FFPE samples when carbamylation was included. Therefore, when further studying protein deregulation and phosphopeptides, carbamylation was excluded for OCT samples, as the results for were not conclusive, but included for FFPE samples.

Comparative MS-based proteomics of OCT and FFPE samples


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By using the optimized protocols, a comparative study of protein expression of two different stages of bladder cancer was performed. Samples of non-muscle invasive (T1) and muscle invasive (T3) tumours, three in each group, were processed as duplicates (Figure 1). For each patient studied, both OCT/frozen and FFPE samples were available (in total 24 samples). On average 1456 ± 167 and 877 ± 125 (average ± SD) proteins were identified in OCT samples and FFPE samples, respectively (Table S1). The overall reproducibility in terms of RSD was 3 and 7% for OCT and FFPE samples, respectively. The number of proteins identified varied between the patients, no matter which preservation method was used. In a specific tumour sample, a higher number was always obtained from the OCT samples as compared to the FFPE samples and when comparing the overlap of proteins from different preservation techniques (Figure 3), it was on average 45%. The low percentage of identical proteins in Patient 4 was strongly influenced by the very low protein coverage in the FFPE sample relative to the OCT sample. On the one hand, this indicated that it was easier to extract proteins from OCT samples and/or that the OCT/frozen approach preserves proteins in a better way for MS analysis. On the other hand, the FFPE samples are more advanced to process due to that both removal of embedding material and reversal of the crosslinking was required. At the time of surgery, the tumour contained certain levels of every protein. After sampling, the time until placement in formalin or OCT and subsequent freezing varied among the specimens and for the two preservation methods, which can further explain the variations.

The correlation of protein expression between the two replicates of samples from the same preservation method was then investigated by plotting the log2 LFQ intensities of proteins in both replicates. The observed Pearson’s correlation coefficients (r) were > 0.9 for all 12 pairs, except for one FFPE sample where r equals 0.64 (Supplementary Figures S1 and S2). These results certified high repeatability of the developed protocols.



Next, comprehensive quantitative comparisons of the two cancer stages using the two different data sets of OCT and FFPE samples were performed. Protein extracts from T24 bladder cancer cells were used as a quality control, depicting that the MS-runs variability was lower that 2% (Table S2). A comparable number of protein identifications were observed for the different tumour samples of the same character (OCT and FFPE groups) (Table S1), again proving that the developed method presented in this work is repeatable. The total numbers of identified proteins in OCT and FFPE data sets were 3036 and 2336, respectively (Table S2). Lists of identified and quantified proteins are provided in Tables S4-S6. In order to compare protein abundances between the two preservation methods, we investigated the protein expression of those 412 proteins that were identified and quantified in both OCT and FFPE sample sets (Table S7). There were also unique proteins for both types of samples that could not be included in this correlation analysis (Table S5 and S6). Ideally, protein abundances should be identical regardless the tissue preservation method. However, Pearson’s correlations > 0.56 were observed for all six patient samples investigated and for four of them the correlation was > 0.74 (Figure S3). From a biological perspective, the values show a positive correlation, indicating agreement in protein expression between the two preservation techniques. These common proteins were also used to perform a PCA (Figures 4A and B). This analysis shows that cancer stage T1 samples are closer to each other, and they are separable from cancer stage T3 samples in the FFPE samples (Figure 4B). The corresponding separation for OCT samples is poorer (Figure 4A). Specifically, the sample from Patient 4 (stage T3) is closer to the stage T1 samples, which could be an explanation of the lowest Pearson’s correlation observed with respect to its FFPE counterpart (Figure S3). To identify proteins that differ between the two cancer stages in the two preservation approaches, a regular Student’s t-test and stringent log2 fold change cut-off of 1 were applied and represented using Volcano plots (Figures 4C and D). This analysis showed that the total number of deregulated proteins, ∼ 25-30, was similar for both preservation methods (Table S2). Nevertheless, the identity of these deregulated proteins differed between the two preservation methods (no overlap). This could be due to the


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heterogeneity of tumours. Even though one and the same tumour was preserved by these two methods, different parts of the tumour had to be directed into the different preservation approaches. The number of samples investigated is also limited and future studies of larger sets will be needed to reveal the full picture. The differences achieved from the different preservation methods underlined that it is important to have access to samples from different patient groups in these investigations, and that the type of preservation plays a crucial role in what proteins are identified and found deregulated. The aim of this study was the development and optimization of methodologies to study preserved tissue by MS-based proteomics. It is noteworthy though, that some of the deregulated proteins in both OCT and FFPE samples were previously established or known as potential cancer biomarkers. For instance, a significant increase in protein expression (4.51 log2 fold change) was detected for transferrin receptor protein 1 (TRFC) in the OCT/frozen invasive tumours, which has also been reported for developing invasiveness in prostate cancer

26,27

. In addition, developmentally-regulated GTP-binding protein 1

(DRG1), up-regulated with a log2 fold change of 2, has previously been localized in bladder cancer tumours. In regard to protein regulation of the FFPE specimens, increased protein expression levels were observed for collagen alpha-1(I) chain (COL1A1), vimentin (VIM), plastin-2 (LCP1), annexin A5, as well as alpha-actinin-1 (ACTN1). These proteins play crucial roles in bladder cancer progression and invasiveness as in agreement with a previous study 28.

Phosphorylation experiments An investigation of protein phosphorylation sites in biobanked tissues was performed. Without enrichment, approximately 20 phosphorylation sites were observed in each sample (Table S2). Of these phosphorylated proteins, 13% overlapped between the OCT and the FFPE samples. As one example, the phosphopeptides in the cancer-related protein HSPB1 (Heat shock 27 kDa protein 1) had a localization score of 100% for the phospho group in both OCT and FFPE samples. Initial enrichments with titanium 19 ACS Paragon Plus Environment


dioxide used 30 µg of protein digest as starting material for three samples (two OCT and one FFPE). The number of identified phosphorylated peptides varied, from 75 in the FFPE sample to 119 in one of the OCT samples (Table S3). The yield, i.e. the ratio of the number of phosphorylated peptides and the total number of peptides identified, was higher than 86%, showing high enrichment efficiency. To maximize the phosphopeptide experiments, we pooled extracts from replicates A and B (originating from one of the patients). Thus, 200 µg protein digest was possible to extract from each type of the samples and four portions of 50 µg were separately enriched with TiO2. The eluted phosphopeptides were subsequently pooled and analysed with LC-MS/MS in three replicate runs. To extract 200 µg proteins from 10 pooled tumour tissue sections of 10 µm thickness from one patient was challenging and not all specimens could provide this much material. For the OCT sample, 211 ± 21 (average ± SD) specific phosphopeptides with unique sequence were identified with a yield of 65%. For the FFPE sample, the corresponding numbers were 133 ± 7 phosphopeptides and 65% yield (Table S3). Compared to non-enriched samples (Table S2) more than 5 times more phosphopeptides were identified. To summarize, provided enough material, several hundred of phosphopeptides could be identified from tissue sections from a single patient. These findings are in good agreement with a statement that FFPE samples can be investigated from the qualitative and quantitative perspective at the PTM level 29. This is, to the authors’ knowledge the first time phosphorylation sites in OCT/frozen samples have been presented. Even though phosphorylation studies of cell extracts using similar amount of starting material report thousands of phosphopeptides30, the numbers of phosphopeptides reported here are reasonable. This was an initial testing of phosphopeptide analysis in our preserved material and the numbers of phosphopeptides identified are likely to increase by further optimization of binding conditions to enrichment material, improvement of enrichment materials and the compositions of binding and elution buffer in enrichment. Also, phosphorylation analysis was not planned at the time for sampling and storage. Very quick preservation to stop all phosphatase activity should be considered and/or adding phosphatase


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inhibitors in some way already at sampling. Conclusions on differences in phosphopeptide patterns can therefore not be drawn from this pilot study.



Conclusions In this study we have demonstrated a rapid, highly reproducible and high-throughput proteomic workflow for subsequent LC-MS/MS analysis of both FFPE and OCT-embedded and frozen clinical tissue samples. The optimized process of removing embedding material for the different preservation techniques followed by efficient protein extraction using urea/thiourea buffer and MED-FASP digestion allows for comprehensive protein identification and relative quantitation of protein expression in FFPE and OCT/frozen specimens. The observed differences of protein profiles from FFPE and OCT should be considered. Our findings also demonstrate the detectability of site-specific PTMs, such as phosphorylation, in both types of biobanked tissues. Moreover, by applying the titanium dioxide enrichment strategy described in this study, a substantially larger number of phosphorylated peptides could be detected in a single pair of FFPE and OCT/frozen samples in comparison with non-enriched material. Although the primary goal of this study was not to discover new diagnostic biomarkers useful for personalized medicine, our straightforward workflow shows great potential to be used for retrospective biomarker discovery studies.

Acknowledgments This work was supported by Åke Wiberg Foundation (SBL, M14-0127), Magnus Bergvall Foundation (SBL, 2015-01200, 2016-01675), Carl Trygger Foundation (SBL, CST 15:57), Clas Groschinsky memory foundation (SBL, M1603, M1742). The authors like to thank Alexander Falk and Ganna Shevchenko for fruitful discussions.

Conflict of interest The authors have declared no conflicts of interest.


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Figure legends Figure 1. Schematic representation of the workflow. Developed workflows for processing of biobanked samples were used to study differences in preservation methods (OCT/frozen and FFPE samples) as well as differences in protein expressions between two stages of bladder cancer. Three samples from each tumour stage were investigated for each preservation method in duplicates (Patients 1-3 were diagnosed with stage T1 and patients 4-6 were diagnosed with stage T3). Relative protein quantifications were used for data comparison. The modification of phosphorylation was studied both with and without enrichment with titanium dioxide enrichments. Figure 2. Total ion chromatograms for the LC-MS/MS profiles for different samples and sample preparations. Panel A: In-solution digestion of an OCT sample. Panel B: Optimized sample preparation workflow for OCT sample. Panel C: Initial testing of removal of paraffin from FFPE samples using a combination of xylene and ethanol. Paraffin peaks were observed at the end of the chromatogram. Panel D: Optimized sample preparation workflow for FFPE sample. Panel E: Optimized workflow applied to T24 cell line (no sample preservation). Figure 3. Overlap of proteins identified in OCT and FFPE samples from six different patients (Patients 13 were diagnosed with stage T1 and patients 4-6 were diagnosed with stage T3). Figure 4. Comparison of label free intensities or expression ratios of proteins commonly found among the two tumour stages samples preserved by OCT-frozen (A and C) or FFPE (B and D) methods (Patients 1-3 were diagnosed with stage T1 and patients 4-6 were diagnosed with stage T3). A) PCA of data from OCT samples; B) PCA of data from FFPE samples; C) Volcano plot of data from OCT samples; D) Volcano plot of data from FFPE samples.



Supporting information:

Figure S-1

Correlation of biological replicates of OCT/frozen samples.

Figure S-2

Correlation of biological replicates of FFPE samples.

Figure S-3

Correlation of FFPE and OCT intensities.

Table S-1

Identified proteins obtained from two biological replicates preserved using

different methods. Table S-2

Identified and quantified proteins obtained from different samples using the

optimized extraction/digestion method. Table S-3

Number of peptides and phosphopeptides obtained using different starting

material. Table S-4

ProteinGroups of OCT and FFPE samples.

Table S-5

Unique and deregulated proteins at different stages of OCT samples.

Table S-6

Unique and deregulated proteins at different stages of FFPE samples.

Table S-7

Common proteins at different stages of OCT and FFPE samples.


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(30) Palmisano, G.; Parker, B. L.; Engholm-Keller, K.; Lendal, S. E.; Kulej, K.; Schulz, M.; Schwammle, V.; Graham, M. E.; Saxtorph, H.; Cordwell, S. J.; Larsen, M. R. Mol. Cell. Proteomics 2012, 11, 1191-1202.



Figure 1


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Figure 2



Figure 3


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Figure

4



82x42mm (230 x 230 DPI)

ACS Paragon Plus Environment

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A Parallel Proteomic Workflow for Mass Spectrometric Analysis of

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