Matrix-assisted laser desorption/ionization imaging protocol for in situ characterization of tryptic peptide identity and distribution in formalin-fixed tissue


Correspondence to: P. Hoffmann, Adelaide Proteomics Centre, University of Adelaide, Gate 8 Victoria Drive, Adelaide, SA, Australia 5005.




Matrix-assisted laser desorption/ionization (MALDI) imaging mass spectrometry provides the means to map the in situ distribution of tryptic peptides in formalin-fixed clinical tissue samples. The ability to analyze clinical samples is of great importance to further developments in the imaging field. However, there is a requirement in this field of research for additional methods describing the characterization of tryptic peptides by MALDI imaging.


This protocol gives highly detailed instructions, with examples, for (1) successfully performing tryptic peptide MALDI imaging on formalin-fixed cancer tissue using a MALDI-TOF/TOF MS instrument, (2) tentatively generating identifications through nLC/MS/MS, and (3) validating these identifications by in situ MS/MS of peptides of interest.


This protocol provides a detailed and straightforward description of the methods required for groups new to MALDI imaging to begin analysis of formalin-fixed clinical samples. Copyright © 2013 John Wiley & Sons, Ltd.


Matrix-assisted laser desorption/ionization imaging mass spectrometry (MALDI-IMS)[1] directly measures the in situ distribution of analytes such as drugs,[2, 3] lipids,[4, 5] peptides,[1, 6-9] and proteins.[1, 10-12] The advantages of the MALDI-IMS approach include the ability to analyze hundreds of analytes in a single experiment without prior knowledge of tissue composition, or the use of antibodies.[13, 14] Furthermore, MALDI-IMS can utilize as little as one tissue section per analysis, making it ideal for precious samples.[8, 15]

To date, the most clinically relevant application of MALDI-IMS has been the acquisition of in situ molecular profiles which can be used to classify tissue types and thereby provide patient specific information.[11, 16-21] Until now, most of those applications utilize frozen tissue, which is less than ideal given that the establishment and maintenance of large frozen tissue collections is challenging and expensive.[22]

Clinical samples for histological studies are traditionally treated with fixatives to prevent the tissue from degrading and to maintain cellular structure.[23] The most common fixative used worldwide in hospitals and pathology institutes for light microscopy is 10% neutral buffered formalin.[22, 24] Formalin (methanol-stabilized formaldehyde) creates intra- and inter-protein cross-links between multiple amino acids, thereby halting tissue degradation and allowing storage at room temperature for decades.[24-26] The longevity of FFPE tissue has led to the accumulation of substantial patient sample archives, which have been collected alongside clinico-pathological data.[27] With suitable methods for accessing FFPE tissues, these archives represent a rich resource for tissue characterization using MALDI-IMS.[26]

In order to circumvent the formalin-induced cross-links most groups have either adopted a proteolytic enzyme[7, 28] or antigen retrieval and enzyme combination treatment.[6, 14] Recently, applied methods demonstrating the usage of antigen retrieval in combination with tryptic proteolytic digestion have shown marked success in reproducibly generating peptides which are discriminatory for known histological features.[6, 29] The methods necessary to prepare FFPE samples, perform MALDI-IMS, analyze data and characterize the resulting peptide features have not yet been standardized and are largely developed in-house by individual laboratories.[6, 7, 14, 30, 31] As such, the need to add to the limited number of detailed protocols in the field[14] prompted the current manuscript, in which the methods developed and published in two previous manuscripts by our group[29, 32] have been described in a single detailed experimental workflow combining tryptic peptide MALDI-IMS and subsequent liquid chromatographic/tandem mass spectrometric (LC/MS/MS) identification of the peptides which localize to a region of interest (e.g. tumour tissue). As such, the current manuscript provides detailed descriptions of the sample preparation, acquisition and data analysis methods needed to repeat the aforementioned result.[32] In addition, key explanations, method expansions and a bio-informatics tool have been provided to make the protocol (Fig. 1) as straightforward as possible.

Figure 1.

Protocol for mapping the distribution and determining the identity of tryptic peptides generated in situ. Following sectioning, tissue is treated with antigen retrieval, digested with trypsin and coated with internal calibrants and matrix. Data is then acquired using a MALDI-TOF MS instrument followed by processing of spectra (smoothing, baseline subtraction, peak picking and internal calibration) and generation of ion intensity maps. Peaks are picked and matched to identifications obtained by nLC/MS/MS analysis of trypsin digested tissue isolated by laser capture micro-dissected tumor tissue from the same patient.[32]


2.1 Chemicals, consumables and equipment

See Table 1 for a comprehensive list of chemicals, consumables and equipment used in this protocol.

Table 1. Comprehensive list of chemicals, consumables, calibrant peptides and equipment required for the protocols outlined in this publication
Chemicals and consumablesCompanyProduct code
α-Cyano-4-hydroycinnamic acid (CHCA)Bruker Daltonics, Bremen, Germany201344
Acetonitrile (ACN) HPLC gradeMerck, Darmstadt, Germany1.00030.2500
Ammonium bicarbonate (NH4HCO3)Merck, Poole, UK103025E – 1 kg
Ammonium phosphate (NH4H2PO4)Sigma-Aldrich, St. Louis, MO, USAA1645-100 g
Citric acid monohydrateSigma-Aldrich, JapanC0706 – 500 g
Ethanol (EtOH) analytical gradeMerck, Victoria, Australia4.10230.2511
Formic acid (FA)Sigma-Aldrich, Steinheim, Germany21909098
Methanol (MeOH) HPLC gradeMerck, Darmstadt, Germany1.06018.2500
Trypsin goldPromega, Madison, WI, USAV5280
Sodium hydroxide (NaOH)Merck, Darmstadt, Germany1.06498.0500
Trifluoroacetic acid (TFA)Merck, Darmstadt, Germany1.08262.0100
Water (H2O, ultrapure, ≥18.2 MΩ from Barnstead purifier)BarnsteadInternational, Dubuque, IA, USAD11951
XyleneMerck, Victoria, Australia1.08681.2500
C18 spin columnsThermo-Fisher Scientific, Rockford, IL, USA89870
Acclaim Pepmap100 15 cm (3 µm, 100 Å)Thermo - Dionex, Amsterdam, The Netherlands164568
Acclaim Pepmap100 2 cm (3 µm, 100 Å)Thermo - Dionex, Amsterdam, The Netherlands164535
HPLC vials (polypropylene)Dionex, Amsterdam, The Netherlands6820.0029
HPLC vial capsDionex, Amsterdam, The Netherlands6820.0028
ImagePrep slide cover slipsBruker Daltonics, Bremen, Germany267942
ImagePrep spray generatorsBruker Daltonics, Bremen, Germany261614
ITO-coated glass slidesBruker Daltonics, Bremen, Germany237001
PEN membrane slides (micro-dissection)MicroDissect, Herborn, Germany11505158
Safe-Lock polypropylene tubes (0.5 mL)Eppendorf, Hamburg, Germany0030 121.023
Safe-Lock polypropylene tubes (1.5 mL)Eppendorf, Hamburg, Germany0030 120.086
Stainless steel cryotome blades (35 × 80 mm)Feather Safety Razor Co., JapanS35
CalibrantSequenceCompanyProduct code
Angiotensin I(34-43)DRVYIHPFHLBioRad, Hercules, CA, USAC10-00002
Dynorphin A(1-17)YGGFLRRIRPKLKWDNQAuspep, Victoria, Australia2032
[Glu1]-Fibrinopeptide BEGVNDNEEGFFSARSigma-Aldrich, St. Louis, MO, USAF3261
Peptide calibration standard IIProprietary peptide mixtureBruker Daltonics, Bremen, Germany222570
EquipmentCompany information
Centrifuge 5810REppendorf, Hamburg, Germany
190 mm polypropylene vacuum desiccatorKartell, Italy
Leica AS LCM microscopeLeica Instruments, Wetzlar, Germany
Microm HM325 microtomeZeiss, Göttingen, Germany
Water bath – Leica HistoBath HI1210Leica Instruments, Nussloch, Germany
LG 700 W MS19496 microwaveLG, China
Proteineer fraction collectorBruker Daltonics, Bremen, Germany
ultrafleXtreme MALDI-TOF/TOFBruker Daltonics, Bremen, Germany
ImagePrep stationBruker Daltonics, Bremen, Germany
MTP 384-800 µm AnchorChip targetBruker Daltonics, Bremen, Germany
MTP slide adaptor IIBruker Daltonics, Bremen, Germany
NanoZoomer automated microscopeHamamatsu, Beijing, China
CanoScan 5600 FCanon, Taiwan
nano-HPLC Ultimate 3000 RS systemDionex, Amsterdam, The Netherlands
Syringe pumpCole-Parmer, Vernon Hills, IL, USA
1/32" PEEK T-pieceUpchurch Scientific, Oak Harbor, WA, USA

Note: Example boxes & figures use data sets published in Gustafsson et al.[32] to demonstrate the anticipated results of the described protocol.

Note: Figures were formatted using R,[33] MS Powerpoint (MS Office 2007), InkScape,[34] and GIMP V2.6.11.[35]

2.2 Protocol-specific solutions

2.2.1 10 mM citric acid monohydrate pH 6[29, 36]

  • 1.05 g citric acid monohydrate (Sigma-Aldrich, Japan)
  • Dissolve in 480 mL H2O
  • pH to 6.0 using 1 M NaOH (~13 mL) (Merck, Darmstadt, Germany)
  • Make up volume to 500 mL with H2O

2.2.2 100 mM ammonium bicarbonate (NH4HCO3) stock

  • 197 mg NH4HCO3 (Merck, Poole, UK) in 25 mL H2O

2.2.3 Internal calibrant solution (100 μL volume with 0.2% TFA v/v)[32]

  • 0.4 pmol/μL angiotensin I34-43 (BioRad, Hercules, CA, USA)
  • 0.4 pmol/μL [Glu1]-fibrinopeptide B (Sigma-Aldrich, St. Louis, MO, USA)
  • 1.8 pmol/μL dynorphin A1-17 (Auspep, Victoria, Australia)
  • 1.6 pmol/μL ACTH1-24 (Sigma-Aldrich, St. Louis, MO, USA)

2.2.4 Matrix solutions (% is v/v)

  • MALDI-IMS (ImagePrep standard solution)
    • 7 mg/mL α-cyano-4-hydroxycinnamic acid (CHCA, Bruker Daltonics, Bremen, Germany) in 50% ACN (Merck, Darmstadt, Germany), 49.8% H2O and 0.2% TFA (Merck, Darmstadt, Germany)[37]
  • LC/MS/MS (Bruker Daltonics method)
    • 748 μL 95% ACN, 4.9% H2O and 0.1% TFA
    • 36 μL saturated CHCA in 90% ACN, 9.9% H2O and 0.1% TFA
    • 8 μL 10% TFA in H2O
    • 8 μL 100 mM NH4H2PO4 (Sigma-Aldrich)

2.3 ImagePrep[38] methods

2.3.1 Trypsin deposition (Bruker Daltonics method)[32]

  • 38% spray power, 0% modulation
  • 30 spray cycles
  • Spray time of 1.25 s
  • Drying time of 45 s

2.3.2 CHCA matrix (adapted from Bruker Daltonics standard CHCA method – see details below)[32]

Phase 1. – Modified to include a minimum of 8 cycles


  • 3.1Prior to beginning, fill the water bath (e.g. Leica HistoBath HI1210, Leica Instruments, Nussloch, Germany) with H2O and set the temperature to 39 °C.
    • If the temperature is set too high the water will melt the paraffin and increase the difficulty of mounting the sections.
  • 3.2Mount the FFPE block onto the Microtome (Microm HM325, Zeiss, Göttingen, Germany) sample holder and ensure the angle of the holder is appropriate. Then insert the microtome blade (35 × 80 mm, Feather Safety Razor Co., Japan).
  • 3.3Set the sectioning thickness to 20 µm and trim the block until the tissue area(s) of interest is visible.
  • 3.4Change the sectioning thickness to between 4 and 6 µm. Begin collecting sections.
    • The thickness which maintains tissue section morphology best and gives the most consistent results should be the standard. Optimum section thickness, in our hands, is 6 µm.
  • 3.5Use a fine-tip brush to lay the tissue section(s) gently onto the surface of the water in the pre-heated water bath.
    • Make sure the sections are not folded or rolled prior to floating in the water bath.
  • 3.6Water mount the sections onto an indium tin oxide (ITO)-coated slide (Bruker Daltonics) by immersing the slide beneath the section (at the desired location) and using it to lift the section out of the water.
    • Mount the section in one smooth motion to prevent section curling or folding.
    • The paraffin edge of the section will stick to the slide. This can be exploited to stabilize the section prior to final section mounting.
    • Finally, ensure that no air bubbles are caught beneath the section. This can cause the tissue section to detach from the slide during antigen retrieval (unpublished observation).
  • 3.7Leave a ~1 cm wide space in the middle of the slide free of tissue. This space is required by the light scatter sensor in the ImagePrep (Bruker Daltonics) sample preparation instrument.
  • 3.8Holding the prepared slide vertically, gently tap excess water onto a lint-free wipe.
  • 3.9Place the slide vertically into a container (e.g. coplin jar or similar) and allow the sections to air dry at room temperature.
    • Optionally, slides can be dried by incubating at 56 °C.
  • 3.10Lay the slide(s) (tissue side facing up) onto a heating block at 60 °C for 1 h.
    • Heating improves the adherence of tissue to the slide.
  • 3.11Wash slides twice in 100% xylene (5 min each, xylene from Merck, Victoria, Australia) to remove paraffin (vertical slide orientation in a suitable glass jar for all washes).
  • 3.12Wash the slides twice with 100% ethanol (EtOH, Merck, Victoria, Australia) (2 min each) and allow the slides to dry.
  • 3.13Mark the slide(s) surface using a water-based white out (e.g. Tipp-Ex) or diamond tipped pen.
    • The marks are used to teach the tissue position in the mass spectrometer.
  • 3.14Place the slide(s) in a coplin jar (or similar) and fill the empty slots with blank microscopy slides.
    • A full jar prevents formation of large bubbles during antigen retrieval and has been observed to reduce section fragmentation (unpublished observation).
  • 3.15Rinse sections twice in 10 mM ammonium bicarbonate (NH4HCO3) for 5 min.
  • 3.16Discard the NH4HCO3 and re-fill the coplin jar with 10 mM citric acid monohydrate at pH 6 (see section 2.2.1)..
  • 3.17With the lid loosely placed on top, put the coplin jar in a microwave and use standard settings (i.e. quick start) to bring the citric acid solution to a boil.
    • The lid should be loosely placed on the coplin jar to prevent a dangerous pressure buildup during microwave heating.
  • 3.18Once the solution reaches boiling point, set the microwave power such that solution temperature is maintained near boiling point. Incubate at this power setting for 10 min.
    • Settings will vary depending on microwave model. Test settings using a set of blank slides first.
    • Our laboratory uses a LG 700 W MS19496 (LG, China) microwave oven.
  • 3.19Following microwave incubation, transfer the coplin jar to a heating block set at 98 °C for 30 min.
  • 3.20Remove the slide(s) following incubation and cool to room temperature.
  • 3.21Rinse the slide using two 1-min dips in 10 mM NH4HCO3 (use fresh NH4HCO3 for the second dip).
    • This step prepares the tissue for tryptic digestion by neutralizing the citric acid remaining on the tissue section. Trypsin activity is optimal at pH 7.5–8.5.
  • 3.22Dry the slide briefly (5–10 min) in a desiccator (no vacuum).
    • Our laboratory uses a 190 mm polypropylene vacuum desiccator (Kartell, Italy).


  • 4.1Scan the slide(s) which will be analyzed at a minimum resolution of 2400 dpi. Our laboratory currently uses a CanoScan 5600 F (Canon, Thailand) scanner.
  • 4.2Ensure that a computer is connected to the ImagePrep for light sensor signal read out.
    • NI VI Logger Lite (V2.0.1, National Instruments, Austin, TX, USA) receives input from the analogue-digital converter attached to the ImagePrep (see ImagePrep user manual, V2.0).
    • Light sensor responsiveness can be tested by blocking light to the sensor. This should reduce the measured signal dramatically.
  • 4.3Perform a single cleaning cycle on the ImagePrep using MeOH (Merck, Darmstadt, Germany) and use lint-free wipes to clean the inside of the main chamber.
  • 4.4Load 200 μL distilled H2O directly onto the spray generator (see ImagePrep user manual, V2.0) and select the trypsin deposition method (see section 2.3.1).
  • 4.5Open the spray offset menu in the ImagePrep other menu and set the Global Power Adjustment such that the spray lasts 30–40 s at 38% spray power with 0% modulation.
    • If the spray does not start during the test, the loaded droplet has probably not reached the spray generator.
    • To fix this, stop the run, open the side door and tilt the solution vial slightly back out of its home position. Tap it back into position. This should move the droplet down onto the spray generator.
  • 4.6Dissolve 100 µg lyophilized trypsin gold (Promega, Madison, WI, USA) in 200 μL 5 mM NH4HCO3.
    • Split the total volume into 40 μL aliquots and freeze these at –80 °C until ready for use.
  • 4.7Mix 40 μL 0.5 µg/μL trypsin gold stock with 160 μL 25 mM NH4HCO3.
  • 4.8Load the full 200 μL of diluted trypsin gold (concentration = 100 ng/μL) onto the ImagePrep spray membrane (see ImagePrep user manual, V2.0).
  • 4.9Start the deposition and monitor the quality of the trypsin spray.
    • The trypsin spray should cover the tissue sections completely and dry (no droplets visible) within the 45 s drying time.
    • The spray should easily reach the end of the slide.
    • The entire volume of trypsin should be deposited over 25–35 spray cycles. Restart the method if required.
  • 4.10Once the entire volume has been deposited, immediately remove the slide from the ImagePrep and place it into an airtight container along with wet paper towels (creates a humid atmosphere).
  • 4.11Place the container at 37 °C for 2 h.
  • 4.12Following completion of the tryptic digest, load the internal calibrant solution (see section 2.2.3) onto the ImagePrep station spray generator.
  • 4.13Place the slide in the ImagePrep station and use the trypsin deposition method to coat the tissue sections with the internal calibrant solution.
  • 4.14While the calibrants are being deposited, prepare ≥5 mL of 7 mg/mL CHCA (see section 2.2.4).
    • At least half of the ImagePrep solution vial needs to be filled to prevent negative pressure in the vial during preparations. Negative pressure reduces spray power (company recommendation).
  • 4.15Change the spray offset, as before, to those indicated in the standard CHCA method (see section 2.3.2).
    • For phase 1 of the matrix method this is 20% power ± 35% modulation.
    • Perform a test spray. The matrix mist should just reach the far end of the ImagePrep and homogeneously cover the central raised area of the ImagePrep chamber.
    • If the spray power is not adequate change the spray offset to compensate.
    • To maintain spray quality, the spray offset may need to be increased in-between phases during sample preparation.
    • Prior to starting the automated preparation wipe away the deposited matrix using a small volume of MeOH and a lint-free wipe.
  • 4.16Place the ITO-coated slide (with experimental sections) onto the central platform of the ImagePrep chamber.
    • The light scatter sensor should not be covered by tissue.
    • Place a glass cover-slip (Bruker Daltonics) over the light sensor (prevents light scatter artifacts interfering with sensor controlled sample preparation).
  • 4.17Begin the matrix sample preparation.
    • Phase 1 is the only preparation phase which is not controlled by the light sensor. As a result, monitor the deposition to ensure that excessive amounts of matrix are not being deposited.
  • 4.18When phase 3 is reached, monitor the preparation using the LoggerVI software to ensure that the automated sample preparation proceeds correctly.
  • The sample preparation from phase 3 onwards relies on formation of a light scatter curve (see Fig. ).
  • If sample preparation is not proceeding correctly you will notice two things:
    1. The spray power boost will increase incrementally (if the spray is too powerful, stop the preparation and restart phase 3).
    2. The cycle number will not change.
  • To correct this you can increase the number of minimum cycles in phase 1 or increase the spray offset for the preparation.
  • The critical step is to achieve a stable light scatter curve by the end of phase 1 (see Fig. ).
  • 4.19Once the preparation is stable in phase 3 it no longer needs to be actively monitored.
  • 4.20When the preparation is complete, remove the slide from the ImagePrep, dispose of the cover-slip and use a small volume of MeOH on a lint-free wipe to wipe away the matrix from a 0.5-cm wide space at both ends of the slide.
  • The space ensures that the metal on the slide adapter contacts the conductive slide surface.
  • 4.21Load the prepared slide into a MTP slide adapter II (Bruker Daltonics).
  • 4.22Re-suspend a tube of peptide calibration standard II (Bruker Daltonics) in 125 μL 0.1% TFA to create a stock solution. Dilute this stock 1:50 with some of the remaining matrix solution (see section 2.2.4).
  • 4.23Sonicate the peptide calibration standard for 5 min and then deposit 0.5 μL of the solution onto the clean slide area left behind by the cover-slip.
  • 4.24Once the peptide calibration standard II spot has crystallized, load the MTP slide adapter into the MALDI-TOF/TOF instrument.
  • Our group uses an ultrafleXtreme MALDI-TOF/TOF instrument (Bruker Daltonics).
Figure 2.

ImagePrep light scatter curve for a single cycle of a typical matrix spray. The light scatter curve shows the original plateau reading, the dip caused by matrix deposition, the subsequent increase of light scatter due to the crystallization of the new matrix layer and the final plateau, which exhibits a higher light scatter than the previous plateau as a result of increased matrix crystal density. The figure was generated in R[33] using log files (ASCII) extracted from the ImagePrep station.


    • NB: This protocol utilizes ultrafleXtreme MALDI-TOF/TOF specific programs and settings.
  • 5.1Load a suitable reflectron positive MALDI-IMS method in flexControl (V3.3, Bruker Daltonics).
  • Settings which can be optimized at this point include:
    • Laser power (instrument specific)
    • Laser offset (instrument specific – typically does not need to be altered for CHCA matrix preparations)
    • Laser repetition rate (typically maximum, 1 kHz for ultrafleXtreme)
    • Detector gain
    • m/z measurement range (at least m/z 1000–3500)
    • Matrix suppression (700 Da, usually 100 Da below m/z range minimum)
    • Acquisition rate (at least 1 GS/s for reflectron MS)
  • 5.2Open flexImaging (V2.1 or newer, Bruker Daltonics) and select create a new sequence in the window that appears. Use the wizard to set up all the relevant acquisition details:
    • Sequence name
    • Data directory
    • Sample preparation type (in this case uniformly distributed coating – 100 µm raster width)
    • AutoXecute method (edited through flexControl)
  • 5.3Use the scanned image from step 4.1 to teach the slide positions on the section image.
  • When the three teach marks have been assigned, select move sample carrier from the edit drop down menu. Move the sample carrier to the edges of the assigned teach marks to confirm accurate teaching.
  • 5.4Move the sample carrier to a region of prepared tissue. Optimize laser power on the tissue to ensure that sample preparation has been successful.
  • Note that this protocol analyzes cellular proteomes. As such any acquired spectra are highly complex and slight arching of the spectrum is typical (see Fig. ) and overlap of peptide isotopic profiles can occur.
  • 5.5An optimum laser power has been achieved when a high intensity spectrum (≥5000 counts for base peak) can be acquired with 500 accumulated shots. The intensity will be variable and can exceed 104 for some spectra.
  • Test the selected laser power on several randomly selected spots.
  • All four internal calibrants should be detectable in a typical high signal-to-noise (S/N) spectrum. Check this prior to starting a full automated acquisition.
  • 5.6Using the same method as steps 5.3 and 5.4, move the sample carrier to the peptide calibration standard II position of the slide. Acquire 500 shots from the calibrant position, ensuring that the peaks in the isotopic profile can be resolved to baseline.
  • 5.7Calibrate the instrument using the standard peptide calibration list provided by the proprietor and save the flexControl method.
  • Use the same or similar laser powers for both the external calibrants and sample spectra acquisitions.
  • 5.8Ensure that the flexControl (V3.3) autoXecute method contains the appropriate parameters.
  • The autoXecute tabs will have the following features:
    • General – flexControl method only selected
    • Laser – set optimum laser power and turn fuzzy control off
    • Evaluation – should contain no background list
    • Accumulation – fuzzy control off, 500 shots acquired in 500 shot steps, dynamic termination off
    • Movement – random walk off
    • Processing – flexAnalysis method required (see below)
    • MS/MS – no method
  • flexAnalysis (V3.3, Bruker Daltonics) method should perform the following processing (see Supplementary Fig. S1 (Supporting Information) for flexAnalysis script required). The following additional settings need to be defined in the flexAnalysis Edit Processing Parameters and Edit Parameters windows.
  • Smoothing (Gaussian, 2 cycles with width of m/z 0.02)
  • Baseline subtraction (TopHat)
  • Peak detection (monoisotopic, e.g. SNAP using averagine peptide composition)
  • Re-calibration(quadratic internal re-calibration, 300 ppm tolerance using a custom mass control list [.mcl] containing the following calibrants):
    • Angiotensin I [M + H]+: 1296.685
    • [Glu1]-Fibrinopeptide B [M + H]+: 1570.677
    • Dynorphin A [M + H]+: 2147.199
    • ACTH1-24 [M + H]+: 2932.588
    • Trypsin (autolysis) [M + H]+: 2211.105
  • 5.9Prior to starting the data acquisition, two important changes to the data-loading settings need to be made. In flexImaging, go to the Edit dropdown menu and select sequence properties. Select read processed spectra, tick the box next to store reduced data for faster access and set the number of data points to 20 000 (only for reflectron data). Select ok. After data acquisition, make sure to save the imaging sequence, which creates a .dat file from the reduced spectra in the results directory.
  • Once the data acquisition is completed, flexImaging will automatically load all the spectral data files.
  • This is not the case if the autoXecute batch runner (Bruker Daltonics) is used to run several imaging sequences. These sequences need to be loaded individually.
  • 5.10Start the imaging sequence from the flexImaging checklist menu or by selecting the start autoXecute run icon.
  • 5.11Once the run has completed, the slide can be removed from the MS instrument and washed in 70% EtOH to remove the matrix (5 min).
  • 5.11Stain the MALDI-IMS section(s) with an appropriate standard protocol. Haematoxylin and eosin (H&E) staining is preferred by our group.
  • Alternatively, a consecutive section can be used for histology.
  • 5.12Following staining, scan the tissue section. Our group uses a NanoZoomer automated slide scanner (Hamamatsu, Japan) to acquire images of stained sections for annotation (20× objective lens).
  • Scans can be annotated by a pathologist to identify the cancerous/non-cancerous region(s).
Figure 3.

Typical MALDI-TOF/TOF spectrum (single) from an in situ acquisition (500 laser shot sum) on an FFPE ovarian cancer section prepared using the described tryptic MALDI-IMS protocol in a previous research article.[32] Note the rising baseline in the m/z range 1000–2000 as well as visual confirmation of three high intensity internal calibrant peaks without zooming (black boxes). All four spiked calibrants in addition to a trypsin autolysis product (m/z 2211.109, used for calibration) should be detected. Inset spectrum shows baseline resolution for angiotensin I and the complexity of the in situ digest. The figure was generated by export of spectral images from flexAnalysis (formatting in MS PowerPoint & InkScape).


    • NB: Although multivariate statistics are best used to select peaks of interest,[12, 41, 42] to keep the workflow as straightforward as possible, peak selection here is manual only and based on operator visualized ion intensity maps for peptides of interest.
    • NB: Steps 6.1 to 6.5 use flexImaging
    • NB: Steps 6.6 onwards use in-house data processing
  • 6.1When the MALDI-IMS data set loads completely in flexImaging (step 5.10), a sum spectrum will be generated in the Spectrum Display window.
  • 6.2Mass filters can be added to this sum spectrum to show the spectral intensity (normalized or non-normalized) within the filter mass range (see Fig. ).
  • 6.3Create a region of interest (ROI) around the tissue section area identified as cancerous (pathologist determined) using the polygon ROI tool.
  • If there is not sufficient contrast in the original section scan to match to the histological stain, co-register the scan of the stained section by selecting Co-Register Image… from the Edit drop down menu in flexImaging. Follow the wizard prompts to complete the co-registration.
  • 6.4In the Regions window select the ROI spectrum checkbox and use the slider at the far right of the Spectrum Display window to move to the corresponding spectrum.
  • 6.5From the cancer-only sum spectrum add mass filters to the monoisotopic peaks of cancer specific peptides (see Fig. ).
  • To add a mass filter while in zoom mode, hold control and click on the sum spectrum at the desired m/z location.
  • To modify mass filter width (absolute or percentage) double click on the mass filter in the Results window list. Typical setting is 0.5 Da.
  • 6.6To generate abundance weight mean (AWM) m/z values for the peptide features in the MALDI-IMS data set, the .xml files (generated by flexAnalysis script) from each individual spectral folder are processed using an in-house developed software tool (available on request from the authors).
    • The software tool (written in Java) uses similar processing work flows to that presented in a previous manuscript.[32]
    • First, .xml peak lists are extracted from the folders containing each individual spectrum.
    • Peak lists are combined into a single list sorted by m/z.
    • Peak groups (i.e. peptides) are defined by applying a linkage distance, e.g. 0.05 m/z units, such that the distance between two adjacent peptide peak groups is always greater than 0.05 m/z.
    • The peaks in each group are then used to calculate an abundance weighted mean.


The larger peptide of interest mentioned, m/z approximately 1905.97 (Fig. 4) was found to have an AWM m/z of 1905.981, as calculated from all spectra using a single linkage distance of 0.05 m/z units.[32]

  • 6.7The AWM m/z values for peaks of interest can now be matched to LC/MS/MS data generated from a tryptic digest of an adjacent section of cancer tissue.
Figure 4.

flexImaging main screen showing the monoisotopic peak selected using a ±0.5 Da width mass filter in the sum spectrum (red outline) as well as an overlay of the H&E stain and ion intensity map produced by the mass filter (a). H&E stain as well as ion intensity maps for two cancer-specific masses (approx. m/z 1628.75 and 1905.97)[32] are shown in (b), (c) and (d), respectively. Ion intensity scales are included. Scale bars are 2 mm (formatting in MS PowerPoint & InkScape).


  • NB: This protocol utilizes laser-capture micro-dissection (LCM) and in-solution treatment of FFPE material (based on methods described in Gustafsson et al.[32]).
  • NB: The following methods refer to the use of an ultrafleXtreme MALDI-TOF/TOF MS instrument as well as the proprietary software required for this instrument.
  • 7.1For LCM, sections of the same FFPE tissue block must be water bath mounted onto PEN membrane slides (MicroDissect, Herborn, Germany).
  • Briefly, FFPE LCM uses a laser system to cut through a polymer membrane the tissue section is mounted on. The free polymer-tissue piece is then collected into a micro-centrifuge tube for further processing.
  • Our laboratory uses a Leica AS LCM (Leica Instruments, Wetzlar, Germany) microscope maintained by a collaborating facility (Adelaide Microscopy, Adelaide, South Australia).
  • 7.2Dry the water bath mounted sections overnight at room temperature.
  • 7.3Heat the PEN membrane slide(s) at 60 °C for 5 min (heating block).
  • 7.4Remove paraffin using a quick 90-s dip in xylene followed by a 60-s wash in 100% EtOH.
  • Avoid longer incubations in xylene as the polymer membrane on the PEN slide will start to degrade.
  • No polymer contamination, attributable to the membrane, has been observed using the protocols described here.
  • 7.5Prepare a solution of 10 mM citric acid monohydrate at pH 6 (see section 2.2.1).
  • 7.6Pipette a small volume of the citric acid solution into the cap of a centrifuge tube suitable for the model of LCM microscope being used.
  • 7.7Load the prepared tubes and slides into the LCM instrument.
  • 7.8Perform LCM and ensure that the pieces have been collected into the cap of the tube.
  • 7.9Following completion of tissue excision and collection, centrifuge the tubes to ensure that all tissue pieces are in the solution at the bottom of the tube.
  • 7.10Add 10 mM citric acid (pH 6) to a volume of 200 μL and heat the tubes for 45 min at 98 °C.
  • 7.11Allow the tubes to cool to room temperature and centrifuge briefly. Remove the citric acid and replace it with 10 μL 25 mM NH4HCO3.
  • 7.12Following a brief incubation (60 s), replace the NH4HCO3 with 10 μL fresh 25 mM NH4HCO3.
  • 7.13Add 10 μL 10 ng/μL trypsin gold and place the digest at 37 °C for 2 h.
  • 7.14Stop the proteolytic digest using 1 μL 10% TFA in H2O.
  • 7.15Use a reversed-phase material to purify the peptides. Our laboratory uses Pierce C18 spin filter columns (Thermo-Fisher Scientific, Rockford, IL, USA) to purify up to 30 µg total peptide. Method based on that provided by Thermo-Fisher Scientific and described in Gustafsson et al.
    • Use one spin column per sample, seated on top of a 1.5-mL micro-centrifuge tube.
    • Centrifuge speed is set to 3000 g for 1 min each (5810R centrifuge, Eppendorf, Hamburg, Germany).
    • To purify peptides, centrifuge the following solutions through the spin columns.
    1. 200 μL 100% ACN (2×)
    2. 200 μL 50% ACN, 49.5% H2O and 0.5% TFA (2×)
    3. 200 μL 2% ACN, 97.5% H2O and 0.5% TFA (3×)
    4. 40 μL 2% TFA in H2O (1×)
    5. 100 μL 2% TFA in H2O (1×)
    6. Replace the micro-centrifuge tubes with a fresh tube.
    7. Centrifuge the peptides through the spin column (×3). Each time pipette flow through onto the column membrane and re-centrifuge.
    8. To wash the peptides, centrifuge through 200 μL 2% ACN and 0.5% TFA (3×).
    9. Replace the micro-centrifuge tube with a fresh 1.5-mL tube.
    10. Elute the peptides using 20 μL 70% ACN and 1% TFA (3×).
    11. Transfer eluate to a HPLC vial and reduce the 60 μL volume to ~5 μL using a vacuum centrifuge.
  • 7.16Increase the volume of the reduced peptide solution to 10 μL using 2% ACN with 0.1% TFA (MALDI) or 0.1% FA (ESI).
  • 7.17The prepared sample can now be loaded onto a capillary or nano-HPLC system connected to an appropriate fraction collector, which can deposit fractions onto a solid sample support (e.g. MTP-384 800 µm AnchorChip target, Bruker Daltonics).
    • Our laboratory uses a nano-HPLC Ultimate 3000 RS Nano/Cap system (Dionex, Amsterdam, The Netherlands) connected to a Proteineer fraction collector (Bruker Daltonics).
    • Method based on that described in Gustafsson et al.[32]
    • Trap column – 2 cm C18 Pepmap100 (3 µm, 100 Å)
    • Analytical column – 15 cm C18 Pepmap100 (3 µm, 100 Å)
    • Solvent A: 97.95% H2O, 2% ACN and 0.05% TFA
    • Solvent B: 80% ACN, 19.96% H2O and 0.04% TFA
    • Inject sample and load for 6 min (5 μL/min) onto trap column at 0%B.
    • Main gradient 8–42%B over 46 min (can be increased to 8–50%B to encompass more hydrophobic peptides)
    • Followed by 42–90%B in 5 min
    • Flow rate set as 300 nL/min
    • Fraction collection every 15 s (184 fractions total)
    • Mix fractions with matrix in a micro-tee junction (PEEK, 1/32" , Upchurch Scientific) prior to discontinuous deposition on MTP-384 800 target (Bruker Daltonics)
    • CHCA matrix (see section 2.2.4) should be supplied at 150 μL/h by a syringe pump (Cole-Parmer, Vernon Hills, IL, USA), combining 625 nL matrix with 75 nL of eluate every 15 s (Bruker Daltonics standard method)
  • 7.18Once all fractions have been collected dilute 1 μL of peptide calibration standard II with 49 μL of 30% ACN with 0.1% TFA.
  • 7.19Further dilute the standard using 150 μL of the CHCA matrix described above.
  • 7.20Spot 0.7 μL of the diluted calibrant onto each solid sample support calibration position.
  • 7.21Open flexControl on the MALDI-TOF/TOF acquisition computer, load the AnchorChip target into the MS instrument and create three autoXecute methods for acquisition of data on the instrument (see Table ).
  • The three methods are for acquisition of calibrant, sample and MS/MS spectra. A general guide to their setup is provided below.
  • When the AnchorChip is loaded, ensure that prior to data acquisition, the source high vacuum has reached at least 1 × 10–6 mbar. Ideally, this value should be below 9 × 10–7 mbar.
  • 7.22The laser power for the individual flexControl methods for each autoXecute method should now be optimized to ensure acquisition of spectra resolved to the baseline with high intensity (ideally ≥5000).
  • Optimize each method on its corresponding spot (i.e. calibrant method on the calibrant spots).
  • Laser power for MS/MS is typically 8–10% more than for MS.
  • 7.23Once you have set up the autoXecute methods, open WARP-LC (V1.2, Bruker Daltonics), select File and new autoXecute run.
  • 7.24Select the appropriate target geometry (e.g. MTP-384 800).
  • 7.25When the wizard prompts, tick the calibration and MS check boxes and select the autoXecute methods for both calibrant and MS sample.
  • These methods need to exist in flexControl before they can be selected here.
  • 7.26Select the data directory, enter the sample name, set the LC delay time (if applicable) and enter the time slice (fraction width in seconds).
  • 7.27Finish the wizard and save the autoXecute sequence in the same directory as the spectra.
  • 7.28Select Start AutoXecute Run (MS or MS/MS) in the WARP-LC main window.
  • 7.29Once the MS acquisition is complete, select Edit for the WARP-LC method line in the main WARP-LC software window.
  • 7.30Save the method as and change the settings in various tabs as indicated in Table .
  • 7.31Select Calculate MALDI compound list. When the calculation is complete, the list can be displayed with the compounds selected for MS/MS indicated by a tick mark.
  • At this point the list can be manually modified to include or exclude peaks of interest, if they are known from previous experiments.
  • 7.32If the list is acceptable (MS/MS on ≥1000 compounds for complex sample), select Extend AutoXecute Run.
  • 7.33Select Start AutoXecute Run (MS or MS/MS).
Table 2. AutoXecute settings for each of the three methods required by WARP-LC
Tab Calibrant AutoXSample AutoXMS/MS AutoX
General Same flexControl method; m/z range: 700–4000; sampling rate of 2 to 4 GS/s; suppress to 500 m/zAppropriately calibrated LIFT flexControl method
Laser Operator determined, fuzzy control offOperator determined (generally +10% power from sample settings), fuzzy control off
Evaluation No background list selectedN/A
Accumulation 2000 shots in 1000 shot steps3000 shots in 3000 shot steps1500 shots in 100 shot steps
Movement Random walk, 200 shots per raster spotRandom walk, 100 shots per raster spot
ProcessingSmoothingGaussian or Savitzky-Golay, 0.01–0.02 m/z for 1 cycleSavitzky-Golay, 0.15 m/z over 4 cycles
Baseline subtractionTopHatTopHat
Peak pickingSNAP, averagineSNAP, averagine
MS/MS N/AN/AWARP feedback selected
Table 3. WARP-LC method used to calculate a compound list for subsequent MS/MS[32]
WorkflowLC/MALDI workflow selected.
AcquisitionMS/MS autoXecute method from Table 2 selected.
S/N threshold – 15; most intense peak within 5 m/z selected for MS/MS; retention time tolerance – 4 fractions; maximum MS/MS acquisitions per spot – 20.
CompoundsMS tolerance – 50 ppm; merge compounds separated by less than seven fractions; background defined as compounds/peaks in ≥20% of spectra.
Result ProcessingNo methods selected – use ProteinScape (V2.1, described in text).


  • 8.1Open ProteinScape (V2.1, Bruker Daltonics) and select New Project from the File drop down menu.
  • 8.2Name the new project, select Finish and right click on the project in the Project Navigator window. Select New Sample
  • 8.3Fill in the sample details and select Finish.
  • 8.4In the WARP-LC main screen select the ProteinScape button in the Send data to… section.
  • 8.5Select the project and sample for the data to be loaded into and select Ok.
  • 8.6Once your data has loaded, a list of MS/MS spectra will appear in the Project navigator window (Fig. (a) – compounds). A 2D view of the LC/MS run (m/z versus retention time, LC/MS survey view) is also provided in the bottom left corner of the ProteinScape interface (Fig. (b)).
  • The three levels shown in Fig. (a) are the project name, sample name and data set name.
  • 8.7Right click on the data set name in the Project navigator window and select Protein search
  • 8.8Save a method as and select the appropriate search parameters for your sample.
  • For example:
    • Search engine: MASCOT[44]
    • Database/Taxonomy: Homo sapiens
    • Enzyme: Trypsin
    • Maximum missed cleavages: 3
    • Global modifications: None
    • Variable modifications: Oxidation (M)
    • Peptide (MS) tolerance: 100 ppm
    • Fragmentation type: LID/CID
    • MS/MS tolerance: 1.0 Da
    • Peptide charge: Instrument dependent (+1 for MALDI)
  • 8.9Select Start to submit the data and search parameters to MASCOT.
  • Under the data set in Fig. (a) there is a search result, MS spectra and the number of compounds which underwent MS/MS.
  • In Fig. (a) the compounds from a loaded .mgf (mascot generic file) file are shown (6184 in total).
  • 8.10Click on the protein search listing which appears (after about 30 s) under the data name in the Project navigator pane of ProteinScape. The listing has an icon that looks like two red bars superimposed on a mass spectrum (Fig. (a)).
  • 8.11The main view tab will change to show a protein list (see Fig. (c)). Click on a protein in the list and the table underneath will populate with the identified peptides from the protein. The LC/MS survey view will also update to show the locations of the identified peptides in the LC run (Fig. (b)).
  • 8.12With a protein selected, hit Ctrl-A to select all proteins. The peptide list will update to show all peptides identified (see Fig. (d)).
  • 8.13Scroll through the peptide list to find your peptide m/z of interest. Check the match error to ensure that it is within ± 20 ppm.
    display math
  • 8.14Any matches made are now considered tentative, and need to be confirmed either by:
  • Immuno-histochemistry
  • In situ MS/MS (see next section)


The peptide of interest mentioned above – AWM m/z 1905.981 – was matched to heat shock protein beta 1[32] which is shown in more detail in Fig. 6. The ppm error between the measured and theoretical (1905.992, ProteinProspector MS-digest[45]) is 5.77 ppm.

Figure 5.

ProteinScape result panels showing the Project Navigator window (a), the LC/MS survey view of all the identified peptides (b, square box indicates positive database hit), the proteins (c) and complete list of individual peptides with positive database hits (d). Peptide m/z 1905.982 is highlighted in (d) as it was a cancer-specific peptide of interest in this example data set which has been described previously[32] (formatting in MS PowerPoint & InkScape).

Figure 6.

In situ MS/MS experiment was performed previously such that a small region of interest on a consecutive FFPE section (from the same patient block) was analyzed by tryptic peptide MALDI-IMS (a).[32] This region was known in these previous results to contain m/z 1905.982 (heat shock protein beta 1 peptide).[32] The resulting MS spectra (b) from the measured region (blue) and a cancer region (red) confirmed the presence of the peptide, which showed a consistent, cancer-specific, ion intensity distribution (c) as compared to full tissue MALDI-IMS data. Comparison of matching in situ MS/MS (d) and LC/MS/MS (e) fragments (b & y ions) for this peptide m/z confirmed the MALDI-IMS to LC/MS/MS match – heat shock protein beta 1 peptide with sequence LATQSNEITIPVTFESR (f). The figure was formatted in MS PowerPoint (2007) & InkScape following spectral export from flexAnalysis.


    • NB: Two options are available – if the original sample used for MALDI-IMS is less than 2 days old, then in situ MS/MS can be attempted directly from this section. If not, then the sample preparation should be repeated for a consecutive section to maximize chances of a successful acquisition. The protocol below outlines in situ MS/MS following a repeat MALDI-IMS measurement of a small tissue region known to contain the peptide of interest (e.g. 1905.981).
  • 9.1Repeat sections 3 through 5 of this protocol using an FFPE section from the same patient block.
  • 9.2Once the measurement has completed and the data has been loaded in flexImaging, use the ROI tool to outline the region containing the peptide of interest, in this case the cancer region (see Fig. (a)).
  • 9.3In the flexImaging interface tick the checkbox next to the ROI that was just created (in the Regions window).
  • 9.4Right click the sum spectrum and change the display type to 2D All Scans. Zoom in on the m/z range of interest and confirm the presence of the peptide(s) of interest (see Fig. (b)).
  • 9.5Apply a 0.5 Da mass filter to the monoisotopic peak of the peptide and confirm the distribution is as expected (see Fig. (c)).
  • 9.6Once the presence and distribution are confirmed, select the Show Single Spectrum tool in flexImaging and select a spectrum from the ROI which has relatively high intensity in the sum spectrum window and overlaps well with the sum spectrum isotopic peptide profile.
  • 9.7Right click on this tissue location and select Open Spectra in flexAnalysis.
  • 9.8In flexAnalysis, select Internal… in the Calibrate drop down menu. When the Internal Mass Calibration window appears, select the mass control list (.mcl) that contains the four internal calibrant peaks and one trypsin autolysis product (see section 5.8).
  • 9.9Set the User Defined Peak Assignment Tolerance to 300 ppm and the Mode to Quadratic. Click Automatic Assign and set the Zoom Range to ± 0.5%. Click on each calibrant individually to zoom in and confirm correct assignment for all five peptides. If all five cannot be found, select another spectrum from the same region.
  • 9.10Once calibration is confirmed, click Calibrate.
  • 9.11In the flexAnalysis Mass List find the peptide(s) of interest and right click. Select Add to MS/MS List. Right click again and select MS/MS List… to open the list. From the MS/MS list window, select Send to flexControl.
  • 9.12In flexControl, select a suitable LIFT method (see Table ), then click the LIFT tab and select the peptide of interest from the MS/MS List drop down list. Once selected the instrument will automatically assign a Pre-Cursor Ion Selector (PCIS) Window Range.
  • 9.13Return to flexImaging and move the MALDI-TOF/TOF sample carrier (as in steps 5.3 and 5.4) to the locations on the tissue in which the peptide(s) of interest exists in high abundance.
  • NB: Use the non-normalized ion intensity maps for this as these provide the absolute peptide signal at all tissue locations.
  • 9.14Once the sample carrier has moved to the selected location, collect between 3000 and 6000 counts for the precursor ion.
  • It is important to zoom in on the precursor ion and ensure that the profiles from multiple acquisitions overlap and are resolved almost to the baseline.
  • The sample carrier may need to be moved several times, as single locations will become exhausted of sample much faster than during LC/MALDI.
  • 9.15Once the precursor ion has been measured, select Fragments in the LIFT tab. The laser power will automatically be adjusted by the software. Set the laser power to between 5 and 10% higher than the laser power required for the parent mass.
  • 9.16Begin measuring precursor ion fragmentation spectra, increasing laser power, as required, and moving the sample carrier to adjacent areas of high peptide signal.
  • 9.17Upon sufficient fragmentation, select Save As… from the panel just below the MALDI CCTV display. In the window that appears, select a default LIFT processing method from the drop down list and tick the Open in flexAnalysis check box. Select a suitable data path and spectrum name (e.g. 1905_LIFT) and click Save.
  • 9.18Once the MS/MS spectrum is open in flexAnalysis and the processing script has finished, select ProteinScape from the Tools drop down menu.
  • NB: Spectrum can also be sent directly to BioTools.
  • 9.19In the ProteinScape window which appears, select Send selected Spectrum. A separate window will appear. Select the Export MALDI target tab and select the relevant project and sample name. Enter a name for the target and select Ok.


The peptide of interest mentioned above – AWM m/z 1905.981 – could be matched to heat shock protein beta 1 using both MASCOT database searches and by manual assignment of the sequence using BioTools Sequence Editor.[32] Figures 6(d) and 6(e) compare the matching in situ MS/MS and LC-MALDI MS/MS fragments which were used to assign the peptides identity as heat shock protein beta 1.

  • 9.20To submit a MASCOT search, in ProteinScape, open all the levels of the imported data (found under the selected project and sample). Right click on the CombinedLIFT level and select Protein Search
  • Use the same search parameters as those for the LC-MALDI (step 8.8).
  • 9.21Select Start to start the database search. When complete, the peptide matches, if found, will be displayed.
  • 9.22Select the search result which appears next to the imported data. If a positive match has been found, select Manual Validation in the Info tab of the Main View panel.
  • 9.23Compare the in situ MS/MS (Fig. (d)) and LC/MS/MS (Fig. (e)) spectra to each other to confirm correct matching. If the initial match was correct there should be a high concordance between the ion series observed in both spectra.
  • 9.24If no match is possible the LC/MS/MS sequence can be directly compared to the in situ MS/MS using BioTools.
    • Send the in situ MS/MS spectrum from flexAnalysis to BioTools by selecting BioTools from the Tools drop down menu.
    • Once in BioTools, select the Start Sequence Editor icon.
    • In Sequence Editor, select File and New Sequence.
    • Enter the sequence of the tentatively identified peptide and modifications, if any.
    • Then select the Send to BioTools icon and return to BioTools.
    • The in situ MS/MS spectrum will now be annotated with any theoretical fragments from the entered peptide sequence.
    • Open the LC/MS/MS spectrum of interest in ProteinScape and select Manual Validation as before.
    • Compare the in situ MS/MS and LC/MS/MS in BioTools.
    • A match can be confirmed by the existence of unique MS/MS fragments in both the LC/MS/MS and in situ MS/MS spectra.


The in situ MS/MS to LC/MS/MS match in Figs. 6(d) and 6(e) shows that the peptide identification by matching was correct, in this case, heat shock protein beta 1. Figure 6(f) shows the peptide sequence complete with fragments found in the LC/MS/MS spectrum (b and y ions).[32]


Formalin-fixed tissues present a unique and rich resource for prospective biomarker and tissue classification studies.[6, 26] Thus, much attention is currently focused on the capacity of MALDI-IMS to provide patient-specific and/or cancer-specific information from formalin-fixed archives and, in particular, tissue micro-arrays (TMAs).[42] Dissemination of this detailed protocol, based on previously demonstrated method advancements,[29, 32] will hopefully lead to the formulation of an international standard MALDI-IMS method for clinical cohorts (e.g. TMAs).[47] Furthermore, by application of the presented method, diagnostic or prognostic peptides may be identified using a combination of MALDI-TOF/TOF and LC/MS/MS: technologies which are readily available to many proteomic groups globally. Finally, the straightforward nature of the presented protocol ensures that groups new to MALDI-IMS can rapidly begin in situ characterization of formalin-fixed samples by MALDI-IMS. As a result, it will be possible for a growing number of laboratories to take advantage of the capacity to generate tissue-specific molecular data for application to clinical questions.


Martin Oehler and Peter Hoffmann acknowledge the support of the Australian Research Council (LP110100693), Ovarian Cancer Research Foundation, the Government of South Australia and Bioplatforms Australia.