TITLE: In Situ Imaging of N-Glycans by MALDIImaging Mass Spectrometry of Formalin-Fixed Paraffin-Embedded Tissue
AUTHORS: Xiaowei Lu, Richard R. Drake, Thomas W. Powers, Kim Norris-Caneda, Anand S. Mehta, Peggi M. Angel, Sean C. Bendall, Michael Angelo

[DESCRIPTION]
Glycosylation of cell surface, secreted, and circulating proteins is one of the most common types of post-translational modification. These modifications occur most commonly as one of three major classes: N-linked glycosylation on asparagine residues, O-linked glycosylation on serine or threonine residues, or as glycosaminoglycan oligosaccharide polymers on serine. Specifically, for N linked glycans, an endoglycosidase enzyme, peptide N-glycosidase F (PNGase F), cleaves the attached oligosaccharides between the asparagine and first sugar. A method to analyze released N-glycans and map them to specific locations within a tissue is presented here. The PNGase F is applied by solvent sprayer as a molecular layer on frozen or formalin-fixed tissues and all released Nglycans in a given region of tissue are detected using matrix-assisted laser desorption/ionization (MALDI) imaging mass spectrometry (MALDI-IMS). Using the described MALDI-IMS protocol, at least 40 or more individual N-glycans can be mapped to tissue histopathology and extracted for further structural analysis approaches.

[STEPS]
SECTION: Dewax
2. You can either use MIBI staining dewax_protocol or the protocol in Drake's published protocol. Here we adapted Drake's protocol into our oven settings in R139. 
SECTION: Antigen Retrieval (AR)
3. The purpose of antigen retrieval is to breakup the cross linking between the amino side chain of lysines and make the tissue more approachable for PNGase F. The preferred antigen retrieval buffer is the Citraconic buffer (pH3) in Drake's protocol. However, based on our experience, MIBI AR protocol and AmberGen AR protocol are all compatible with PNGase F digestion. 
SECTION: Application of PNGase F
4. This step is to let PNGase F cleave off the N-glycan from tissue on the solid phase. The cleaved off N-glycan will stay at its specific location and hence provide spacial information. Before the incubation step, if you messed up the spraying step, you can wash the slide with HPLC water, dry the slide and spray again. However, after the incubation step, when the N-glycan got cleaved off the tissue, the tissue need to be dry. A drop of water will dissolve the N-glycans and compeletly kill the spacial information. After the PNGase F incubation step, if you took slide (vacuum sealed bag) from the freezer, let the vacuum bag warm up to room temperature and then open the bag. Exposing a cold slide to atmosphere will lead to moisture condensation on the tissue. 
SECTION: Slide scanning
5. Optical image will be used for setting up a MALDI run with flexImaging software. 
SECTION: Application of Matrix
6. 
SECTION: Storage of prepared Slide
7. Prepared slides could be placed in mailer in a vacuum seal bag and store at -20 °C for up to a month before MALDI acquisition. 
SECTION: FFPE tissue slide &amp; storage
1. FFPE tissue slides for MIBI and other epitope based stain protocol are usually stored in vacuum sealed bag with anti oxidant and drying agent. FFPE tissue slides for N-glycan IMS purpose can be stored the same way as MIBI slide. However, slides left on bench top at room temperature exposed to atmosphere over a year can be used for N-glycan IMS as well.

Formalin fixed tissues should be sectioned at 3~7 µm and mounted on slides. ForMALDI_TOF, sections should be mounted onto conductive slides. For MALDI-FT-ICR and MALDI-QTOF, standard microscope slides, positively charged microscope slides or epoxide coated microscope slides are suitable as long as they fit into the 25 x 75 mm slide holder for the instrument. 

If a slide with tissue on it does not fit the sample holder for the instrument, it may be sized to fit while the tissue is still covered with wax, as this prevents contamination and limits damage to the tissue. A small hand held rotary tool may be used to grind down slide edges to fit into the instrument sample holder. Afterwards, slides may be lightly rinsed in tap water and allowed to air dry.
SECTION: Dewax
2.1. Heat slides at 70°C (face up) for one hour, and then at 70°C (perpendicular) for 10 min. 
SECTION: Dewax
2.2. Remove and cool to room temperature, usually 5 minutes.
SECTION: Dewax
2.3. You can do dewax washes the same way as you do for MIBI staining slide. Or you can just use plastic/glass Coplin jars and let the slide sit in the solution. Here are the sequential washing solutions:

- Xylenes 3 minutes, repeating a total of two times.

- 100% ethanol 1 minute, repeating a total of two times.

- 95% ethanol 1 minute

- 70% ethanol 1 minute

- Distilled water 3 minutes, repeating a total of two times.
SECTION: Dewax
2.4. Slides may be stored overnight in vacuum desiccator. Dry slides could be placed in a mailer inside a vacuum sealed bag and stored in freezer overnight.
SECTION: Antigen Retrieval (AR)
3.1. Prepare Citraconic buffer:

- 25 mL distilled water or HPLC grade water into a 50 mL falcon tube.

- Add 25 µL of Citraconic anhydride to the water.

- Add 2 µL of 12 M HCl.

- Agitate tube after capping.

- Add water to a total of 50 mL.

- Agitate tube to mix.

- Check that pH is around 3.0 ±0.5 by spotting 2 µL of the prepared buffer onto a pH strip.
SECTION: Antigen Retrieval (AR)
3.2. Heat slides in vegetable steamer:

- Preheat the vegetable steamer for 5 minutes prior to retrieval.

- Add around ~10 mL of the buffer to a 5 slide mailer.

- Place no more than 3 slides per 5 slide mailer. Slides should be placed with tissue facing outward to the solution in positions 1 and 5, and NOT facing the slide mailer walls. Position 3 may face either way.

- Completely fill the slide mailer the rest of the way with the buffer.

- If the mailer has no holes punched in the lid, only snap close one corner of the mailer.

- Place the mailer in the center of the vegetable steamer.

- Set heating for 30 minutes.
SECTION: Antigen Retrieval (AR)
3.3. Cool the slides after antigen retrieval:

- Remove mailer and place in a tub with cool water from the faucet. Water should not go over the top of the mailer.

- Allow to cool for 5 minutes.

- Remove half the buffer from the mailer and replace with distilled water.

- Allow to cool 5 minutes on countertop.

- Repeat removal of half the buffer two more times, each with 5 minutes of cooling.

- Complete by rinsing in 100% distilled water.

- Dry the slides with Kimwipes (don't touch the tissue). And dry slides at least 5 min in a desiccator, then proceed to the PNGaseF digestion steps. 

SECTION: Application of PNGase F
4.2. Prepare PNGase F solution:

- Prepare 0.1 µg/µl PNGaseF in water using aliquots from the -20°C freezer. The tube contain 100 µg PNGaseF could only take 0.5 mL water. Pipet 0.5 mL water into the tube with 100 µg lyophilized PNGase. Transfer out to a 1mL glass flask or eppendorf tube. Repeat for one more time. 

- Ensure that enough solution is prepared, e.g, four slides takes approximately 1 mL of solution; for one slide, take 0.5 mL solution. Once you select the spraying area, HTX sprayer will give you an estimate. Please add 0.25 mL dead volume to that estimate volume. 
SECTION: Application of PNGase F
4.3. Prepare the HTX sprayer:
Bangelo lab is using HTX M3+ Sprayer, It is slightly different from HTX M5 sprayer which was used in Drake protocol. But the spraying method and conditions are exactly the same. 

- Open the nitrogen gas from the nitrogen tank. Switch on the HTX sprayer. Then open the HTX sprayer software. Please follow this exact sequence. Adjust Nitrogen to ~5 psi by turning the nob on the sprayer. 

- Switch on the hood where the HTX sprayer locate.

- Check the level of the four solvent bottle and make sure the tubing are inside the solvents. HPLC water need to be freshly filled within 7 days. 

- Placed the waste solvent beaker under the waste solvent tube. 

- On HTX sprayer software interface, click "setting" (on the bottom of the left column)-->click "pump scripts" --> click through the buttons on the first line to completely clean up the sprayer system.

  

SECTION: Application of PNGase F
4.1. Humidity Chamber Preparation:

- Obtain a 100 x 15 mm cell culture dish with lid.

- Cut a paper towel (Wypex60 preferred) to fit to the bottom of the cell culture dish and place in bottom of dish.

- Take an 11x21 mm Kimwipe (Kimtech) and fold it in half along the creased line. Roll up the Kimwipe lengthwise forming a roll about 10 mm in diameter.

- Place the rolled Kimwipe on one end of the paper towel. You may need to squirt a little DI water on it to make sure it stays rolled.

- Place a second rolled Kimwipe at the other end.

- Saturate the paper towel and Kimwipes with water. Typically, this takes around 5 mL DI water. To perform this, simply hold the incubation dish at an angle and squirt DI water onto the towels and Kimwipes until water starts seeping out from under the towel, indicating saturation. Discard excess water that is not absorbed by the towels and Kimwipe.

- Pre-warm the incubation dish in a 38.5 ± 1.5°C oven at least half an hour. Light condensation should be observable on the lid of the dish. Don't stack up petri dishes inside an oven. 
SECTION: Application of PNGase F
4.4. Spray a dummy slide with HPLC water:

- Place the tubing (labelled as "Enzyme") of port E into a Eppendorf tube with 1 mL HPLC water.

- On HTX sprayer software interface, click "method" (left column) --> choose "PNGase_00" --> double confirm the parameters: speed (25 µL/min), nozzle temperature (45°C), 15 passes, velocity of 1200, 3 mm offset, 10 psi, dry time (zero). 

- Click on "Open Tray Drawer" -->  tape the dummy slide into the sprayer. --> click on "Close Tray drawer"

- Click on "Sample" --> drag the red rectangle to define the spraying area. --> click on "confirm area"

- Click on "Start" and follow the instructions on the HTX sprayer software. 

- when the spray start, pay attention to two things: 

1st, is there a consistent tail made from colorless fog on the slide? Does the length of the tail make sense? If the tail is too long or the liquid does not got evaporated efficiently, something is wrong. 

2nd, when the spraying finished, there should be no trace of salt or any other solid on the dummy slide. The presence of solid on the dummy slide indicate the sprayer system need to be further cleaned. 
SECTION: Application of PNGase F
4.5. Spray PNGase F on tissue slide

- After dummy slide run finished, click on "repeat", if you want to spray exact the same area as dummy slide; click on "new run", if you will change the spraying area. You don't need to click "wash" to proceed with washing steps. 

- Place the tubing (labelled as "Enzyme") of port E into the container with PNGase F solution.

- Click on "Open Tray Drawer" -->  tape the tissue slide into the sprayer. --> click on "Close Tray drawer"

- Click on "Sample" --> drag the red rectangle to define the spraying area. --> click on "confirm area"

- Click "start" and follow the instructions form the sprayer. 

- When the run finished, take out your slides first, adjust the Nitrogen pressure to 5 psi, and click "wash"

- Wait until the washing finished. Close the HTX sprayer software, switch off the sprayer and turn off the nitrogen flow from the nitrogen tank. Please follow this exact sequence. 

- Pour the waste solvent from waste beaker to waste bottle and cap the bottle. Switch off the hood. 
SECTION: Application of PNGase F
4.6. - Place tissue slide inside the incubation chamber (face up). Place the incubation chamber back to the oven and incubate for 2 hours.

- When removing, be aware that heavy condensation will have developed underneath the slide. Remove the slide slowly while holding it parallel with the countertop. Wipe off the condensation before rotating the slide.

- Store the slide in a 5 slide mailer to protect the released glycans. If matrix cannot be sprayed the same day, store at -20°C (in vacuum sealed bag). It is preferred to immediately spray matrix onto the slide.
SECTION: Slide scanning
5.1. Mark fiducials at the four corners of each slide.

Fiducials are points of reference used when “teaching” the instrument software (flexImaging) the coordinates of the slide. Fiducials allow accurate targeting of tissue regions for imaging.

As shown in Figure 2, use a reflective metallic marker to make a small circle at each corner. Use a black marker to draw a cross or hash mark on top of each silver circle; the reflective marker provides a contrasting background for easy visualization of the black fiducial mark by the instrument camera.
  

SECTION: Slide scanning
5.2. Scan the slide following Bruker Scanner manual
 
- placed the slide (face up) on the film of the scanner (connect to processing computer#1). Adjust the level of the film with the four buttons (red square).Close the scanner lid.
 
- Switch on the scanner 

- open the scanner software   
- In the EPSON Scan software, make sure you have applied the following settings:
Mode: Professional Mode
Document type: Film (with Film Holder)
Image Type: 24-bit Color
Resolution: 3200 dpi
Scale: 100%
  - Click the “Preview” button on the bottom left

- Hold down the left mouse button and drag over the slide in the Preview window. And make a dotted line box will form around the low-resolution scan of your slide. This dotted box defines the region of interest (ROI) for the high-resolution scan.
  - Click on "Histogram Adjustment"

 - Adjust the numbers in red rectangle box until you can see the tissue and fiducials clearly. Then click "done".
  - Click the "Scan" button and A file save setting dialog box opens. Save the file on local D drive of processing computer #1: (D:)>Scan_processing computer
 - Open image file with "Paint" --> flip and rotate the image so that it will be the same orientation as it is in the MALDI slide holder. 

SECTION: Application of Matrix
6.1. Prepare the CHCA matrix:

- Prepare CHCA matrix (sealed with para film and stored in freezer) at 7 mg/mL in 50% acetonitrile/0.1% TFA.Add 0.042g CHCA to 6ml 50% acetonitrile/0.1% TFA. Prepare fresh each time in a 50 mL falcon tube.

- Vortex violently (5~10 min) until most solid dissolve.

- Filter CHCA solution using Millex (Millipore) 0.2um syringe filter.  

Small chunks may remain in the bottom of the falcon tube. Make sure that these are not loaded into the TM sprayer loop as they will clog components of the TMsprayer. After the filtration, cap the flask.   Crystal may appear if the solvent evaporated. 
SECTION: Application of Matrix
6.3. Spray a dummy slide with CHCA solution

- Place the tubing (labelled as "Matrix") of port D into the matrix solution.

- On HTX sprayer software interface, click "Method" (left column) --> choose "CHCA_dry" --> double confirm the parameters: concentration (7 mg/mL), solvent (50%ACN, 0.1%TFA), nozzle temperature (77°C), pressure (10 psi), flow rate (70 µL/min), number of passes (14), velocity (1300), Track space (3 mm) , dry time (0), pattern (cc).

- Click on "Open Tray Drawer" -->  tape the dummy slide into the sprayer. --> click on "Close Tray drawer"

- Click on "Sample" --> drag the red rectangle to define the spraying area. --> click on "confirm area"

- Click on "Start" and follow the instructions on the HTX sprayer software. 

- when the spray start, pay attention to two things: 

1st, is there a consistent tail made from light yellow solid on the slide? Does the length of the tail make sense? If the tail is too long or the liquid does not got evaporated efficiently, something is wrong. 

2nd, when the spraying finished, there should be a fine layer of yellow solid on the slide. 
SECTION: Application of Matrix
6.2. Prepare the HTX spray the same way as in section 4.3
SECTION: Application of Matrix
6.4. Spray CHCA on tissue slide

- After dummy slide run finished, click on "repeat", if you want to spray exact the same area as dummy slide; click on "new run", if you will change the spraying area. You don't need to click "wash" to proceed with washing steps. 

- Click on "Open Tray Drawer" -->  tape the tissue slide into the sprayer. --> click on "Close Tray drawer"

- Click on "Sample" --> drag the red rectangle to define the spraying area. --> click on "confirm area"

- Click "start" and follow the instructions form the sprayer. 

- When the run finished, take out your slides first, adjust the Nitrogen pressure to 5 psi, and click "wash"

- Wait until the washing finished. Close the HTX sprayer software, switch off the sprayer and turn off the nitrogen flow from the nitrogen tank. Please follow this exact sequence. 

- Pour the waste solvent from waste beaker to waste bottle and cap the bottle. Switch off the hood.