Profiling Anti-Neu5Gc IgG in Human Sera with a Sialoglycan Microarray Assay

Human sera samples were obtained from the Israeli Blood Bank and were used in accordance with the Helsinki declaration and Tel Aviv University Institutional Review Board.

1. Array Fabrication Planning and Layout

1. Determine the slide layout.
   NOTE: Each slide contains 16 sub-arrays divided into 16 identical blocks numbered B1 to B16 (Figure 1B, Supplementary Figure 1A).
2. Determine the pin layout. Use four pins, each printing four sub-arrays (blocks) per slide:
   Pin 1 prints Blocks 1, 2, 9, and 10;
   Pin 2 prints Blocks 3, 4, 11, and 12;
   Pin 3 prints Blocks 5, 6, 13, and 14; and
   Pin 4 prints Blocks 7, 8, 15, and 16 (Supplementary Figure 1A).
3. Determine the block and 384-well plate layout.
   NOTE: The order in which the glycans appear in each block requires careful planning. In this example array, print each sample at four replicates, design fluorescent marker spots to be located at the bottom-right corner (to facilitate spot analysis), and print standard-curve IgG (STD) spots at six increasing concentrations (Supplementary Figure 1B).
   1. For each printed spot, first dispense the material into four wells (one for each pin) in a 384-well plate.
      NOTE: The order of the materials in the 384-well plate relies on the designed block layout (Supplementary Figure 1B-C).

2. Preparation of Glycans and the Source Plate

1. Prepare glycan printing buffer (50 mL of 300 mM phosphate buffer, pH 8.4). Weigh 58.5 mg of monosodium phosphate monohydrate (NaH₂PO₄.1H₂O) and 3.9 g of disodium phosphate heptahydrate (Na₂HPO₄.7H₂O) in 40 mL of deionized water (DIW). Titrate to pH 8.4 using 4 M NaOH. Adjust the volume to 50 mL and filter through a 0.2-µm membrane for sterilization.
2. Prepare marker buffer (50 mL of 187 mM phosphate buffer, pH 5). Weigh 289 mg of monosodium phosphate monohydrate (NaH₂PO₄.1H₂O) and 1.94 g of disodium phosphate heptahydrate (Na₂HPO₄.7H₂O) in 40 mL of DIW. Titrate to pH 5 using 1 M HCl. Adjust the volume to 50 mL and filter through a 0.2 µm membrane for sterilization.
3. Prepare STD curve buffer (50 mL of PBS-glycerol: phosphate-buffered saline, pH 7.4, supplemented with 10% glycerol from a 100% glycerol stock) and filter it through a 0.2-µm membrane for sterilization.
4. Prepare each glycan stock solution at 10 mM in DIW. Verify the sialylated glycan concentrations by fluorescence detection on reverse-phase high-pressure liquid chromatography analysis¹⁸.
5. Dilute each glycan to 100 µM in a total volume of 100 µL of glycan printing buffer in microcentrifuge tubes.
6. Prepare primary amine containing fluorescent dye. Solubilize Alexa 555-hydrazide to 1 mg/mL with marker buffer and then dilute to 1 µg/mL in a 1 mL total volume.
   NOTE: Any other primary amine-containing dye can be used with an appropriate wavelength according to the slide scanner.
   NOTE: Marker spots facilitate grid alignment in each block during analysis.
7. Prepare human IgG STD curve dilutions: prepare 0.5 mL of a stock solution of human IgG at 160 ng/µL in STD curve buffer. Serially dilute the stock 1:1 six times to obtain 80, 40, 20, 10, 5, and 2.5 ng/µL, all in STD curve buffer at a total volume of 200 µL each.
   NOTE: Other Ig isotypes can be used if suitable for the experiment (e.g., human IgA, IgM, IgE, or an Ig of a different organism).
8. Place the 384-well plate on ice. Using an electronic multi-pipette, aliquot 7 µL of each glycan, marker, and STD curve IgG-four replicates of each-according to the plate layout (Supplementary Figure 1C). For better accuracy, load 35 µL of each sample and aliquot 7 µL into each of the four wells of the sample. Place the remaining 7 µL of each glycan back into their original tube.
9. Cover the plate with parafilm and spin down for 2 min at 250g and 12 °C. Place the plate at room temperature (RT), protected from light. Do not uncover the plate before the humidity in the room reaches 60-70%.

3. Programing the Nano-printer

1. Open the "Microarray Manager" software. Use the "Method properties" window.
2. Set up the work table layout: choose the "56 slide configuration."
   NOTE: This is a predefined configuration that allows for the printing of 56 slides. Calibrate for each type of plate, pin, and print format (Supplementary Figure 2, step 1).
3. Select "Pin Configuration" | "Pin Tool 1×4 9 mm."
   NOTE: This is a predefined configuration for using four pins (Supplementary Figure 2, step 2).
4. Define the "Pin type." Calculate and create the pin type that is suitable for the printing.
   NOTE: This step defines how many spots a pin would print before re-dipping in the sample 384-well and should be optimized for the different pin types and slide surface chemistry.
   NOTE: When using 946MP3 pins, consider two important issues: 1) each pin can print up to 140 homogenous spots and 2) pre-spotting is essential to drain excess liquid from the tip of the pin before printing on the slides. Therefore, program each pin to print 20 pre-spots on a pre-printing block (optimized for 946MP3 pins on epoxy-coated slides).
   1. Calculate the total number of printed spots per sample.
      NOTE: Here, each pin prints 4 spots per sub-array (block); a total of 4 sub-arrays per slides = 16 spots/slide. After 7 slides, each pin prints 16 x 7 = 112 spots. In addition, 20 pre-spots before printing gives 132 spots/dip. Therefore, the pin type is defined as "946MP3-132 spots" (4 spots per sample per block x 4 blocks x 7 slides + 20 pre-spots = 132).
   2. Create the "Pin type" (Supplementary Figure 2, step 3). Press "Setup" (left panel of the software) | "Micro Spotting Pins" | "New Name" (946MP3-132 spots). Define the number of spots per dip (132), the spot diameter (100 µm), the uptake (0.25 µL), the delivery (0.7 nL), and the minimum spot spacing (100 µm). Press "OK" | "OK."
   3. In the "Method properties" window, select the created pin type: "946MP3-132 spots."
5. Define the washing conditions. Set the "Wash before starting" and "Wash after Finishing" to "Normal wash" (Supplementary Figure 2, step 4).
6. Define the "Plate list."
   NOTE: This is the sampling pattern from the plate (the order of dips from the 384-well plate) according to the plate layout. To upload the plate into the plate list, set up a plate sampling pattern.
   1. Create the plate "Sampling Pattern." Go to "Templates" (Supplementary Figure 3) | "Sampling Patterns." Give a new name (e.g., JoVE array) and select "Insert" | "Fill top-bottom" | "Pin Tool 1 x4 9 mm." Select the order of well pickups according to the pre-defined plate layout (Supplementary Figure 1A). Press "OK" | "OK."
   2. Load the plate sampling pattern. Click the right side of the "Plate list" cell (Supplementary Figure 3) | "Add plate" and select the correct plate sampling pattern (the "JoVE array" described in step 3.6.1.). Select "Normal Wash." Press "OK" | "OK."
7. Go to the "Sample Plates" and change the position offset to 1 (which determines where the plate is located on the arrayer deck).
   NOTE: The offset position of 0 is closer to the sonicator bath.
8. Define the "Microarray Print Design." Calculate the block-to-block distances and spacing.
   1. Consider the dimensions of the developing tool that divides the slide into 16 wells of 7 x 7 mm, with 2-mm spaces between sub-arrays (a total of 9 mm between blocks; Supplementary Figure 4).
   2. Consider the sub-array dimensions.
      NOTE: The spacing between spots is 0.275 mm and there are 12 columns and 18 rows in this print design; the total dimensions are: width (11 x 0.275 mm = 3.025 mm) and height (17 x 0.275 mm = 4.675 mm). Calculate the horizontal spacing: 9 mm – (11 0.275) = 5.97 mm. Calculate the vertical spacing: 9 mm – (17 x 0.275) = 4.325 mm. Calculate distance from the left side of slide: 4.43 mm + 1.07 mm = 5.5 mm. Calculate the distance from the top side of the slide: 2.95 mm + 0.55 mm = 3.5 mm (Supplementary Figure 4).
   3. Define the "Print Designs" (Supplementary Figure 5). Select "Templates" | "Print Designs" | "New."
      1. Select "Pin Setup" | "Pin Tool 1 x 4 9 mm" | "Pin – 946MP3-132 spots." In the "General" tab, change the "Name of Microarray Print" (i.e., "JoVE array"). Select "Spot Speed" | "Replicate Number" (4) | "Touch point Height" (0 mm) | "Soft Touch Height" (3 mm) | "Print to all available positions" (Supplementary Figure 5A).
      2. In the "Microarray" tab, choose the horizontal and vertical spacing (5.9 mm and 4.325 mm from step 3.8.2) and the number of horizontal and vertical microarrays (2). Select "Print Microarray Horizontally" | "Print Duplicate Microarray" (Supplementary Figure 5B).
      3. In the sub-array tab, choose the maximum number of columns (12), the maximum number of rows (18), and the distance between the columns and rows (275 µm) (Supplementary Figure 5C).
      4. In the measurements tab, choose the distance from the left side of the slide (5.5 mm), the distance from the top of the slide (3.5 mm, calculated in step 3.8.2), the width of the print slide (25 mm), and the height of the print slide (75 mm) (Supplementary Figure 5D). Press "OK" | "OK."
9. Click the right side of the "Microarray Print Designs" cell and select the defined design (the "JoVE array" from step 3.8.3; Supplementary Figure 6).
10. Define the "Slide Count" (56), and change the "Position Offset" (0). This determines the position of the first slide to be printed on the arrayer deck (Supplementary Figure 6).
11. Define the "Microarray Pre-Print Design" (Supplementary Figure 6). Select "Glass Block" | "Pre-Print 1 x 4 mm Pin tool."
    NOTE: This spot-spacing setting allows 46,464 pre-spots (for all 4 pins) on the glass block, thus 46,464 / 4 = 11,616 pre-spots per pin. When the space on the pre-print block had been used up, the printer will stop and the block will need to be flipped over.
    NOTE: To calculate after how many samples the pre-print block is full, consider the following: every sample will be taken by 8 pin-dips in the 384-well plate to complete 56 slides (7 slides per dip), and each dip has an additional 20 pre-spots, giving a total of 160 pre-spots per sample. Therefore 11,616 / 160 = 72.6 samples. Thus, the pre-print glass block would need to be flipped over after 72 samples (to evaluate the time for flipping, measure how long it takes for one sample to complete a full run on 56 slides and then multiply by 72).
12. Select "Use PrePrint Design" | "Yes" | "Scan Barcodes" (No) | "Print Single Replicate" (No) (Supplementary Figure 6).
13. Click on "Validate" (Supplementary Figure 6).
14. Click on "Save" and save the method as an arr file (e.g., "JoVE array.arr," Supplementary Figure 6).

4. Printing Designed Arrays

NOTE: All steps should be carried out in a clean room with a humidity of 60-70% and with appropriate gloves and clothing for protection

1. Turn on the nano-printer machine and humidifier.
2. For the wash buffer and humidification, fill with DI water. Empty the wash bottle (drain) and humidification trap carboy (Supplementary Figure 7A).
3. Set the humidifier to 70% humidity and start humidification.
4. Clean all pins before printing. Prepare 50 mL of hot (65 °C) pin-tip cleaning solution (15 g of pin-tip cleaning reagent and 50 mL of 65 °C water, mixed thoroughly until the solid reagent dissolves completely).
   1. Apply the cleaning solution to each pin tip by dipping the pin-tip brush (fine) into the pin-tip cleaning solution and brushing the surface of each pin tip.
      NOTE: Pin-tip cleaning solution is corrosive and toxic. Make sure to wear gloves and safety glasses at all times when using this solution. Soaking pins in pin-tip cleaning solution for more than a few minutes may result in permanent pin damage.
   2. Fill the ultrasonic bath with 1 L of distilled water and place the pins in a floatable pin cleaning rack in the bath. Sonicate for 5 min and remove the pins. Let dry and gently wipe with clean room special wipes.
5. Open the "Microarray Manager" printer software; upon connection set, the printer light will turn on (red/green). Press "Stop" | "Clear" | "Init" (Supplementary Figure 7B, steps 1-3); the print-head arm will then reset the X, Y, Z alignments.
6. To load the pins into the print head, press "Load" (Supplementary Figure 7B, step 4) and place the pins in the print head according to the defined configuration of the print/pin tool layout (Supplementary Figure 7C); this layout uses 4 pins, each printing 4 sub-arrays on each slide to get total of 16 sub-arrays per slide (Supplementary Figure 1A). After placing the pins, press "Init" again.
7. Assemble the slides on the arrayer deck. Open the slide box and clean each slide by blowing ultra-high-purity nitrogen gas. Place the desired number of slides on the array platform. Hold the slides with two fingers at the bottom corners and then place the slide into its position. Hold the two right corners and push the left corners gently until the slide tightly fits into its position. Push gently towards the left-bottom corner to ensure a tight fit (Supplementary Figure 7D).
8. Clean the pre-block glass with distilled water, 70% ethanol, and 100% ethanol and let it air dry (use only dedicated clean-room wipes). Place the pre-block glass into its position on the deck.
9. Fill a sonicator bath (Supplementary Figure 7B). Press on "Fill" for 55 s then press "OK." Drain the sonicator by clicking "Drain" for 20 s and then pressing "OK." Repeat the filling and draining once more and then fill again for 55 s.
10. Fill the washing bath (Supplementary Figure 7B): press "Prime" for 40 s and then press "OK."
11. Position the 384-well plate with aliquoted samples into its location on the arrayer deck and remove the parafilm cover.
12. Start printing.
13. When printing is complete, drain the sonicator bath (Supplementary Figure 7B) and turn off the humidifier.
14. Allow the slides to dry on the arrayer deck for 16-24 h without humidification.
15. Number the slides with an alcohol/solvent-resistant marking pen and store them in a vacuum-sealed box in the dark.

5. Processing and Developing the Arrays

1. Take out a slide from the vacuum-sealed box and place it in a chemical hood for 5 min, array facing up, to evaporate the excess liquid.
2. Scan the slide at the highest gain (950 PMT) to ensure that all the spots have been printed.
3. Proceed with the chemical blocking of epoxy groups on the slide
   1. Prepare 50 mL of ethanolamine blocking solution (0.1 M Tris-HCl, pH 8 and 0.05 M ethanolamine). Mix 140 mL of DIW with 8.8 mL of Tris-HCl (1.7 M, pH 8) and 0.45 mL of ethanolamine (16.6 M). Transfer to a 50-mL staining tube (Supplementary Figure 8A) and warm to 50 °C.
   2. Hydrate the slide. Place the slide in a staining tube filled with DIW, cover it with foil, and incubate for 5 min with slow and gentle shaking at RT.
   3. Block the reactive epoxy groups on the slide. Dip the slide into the 50-mL staining tube filled with pre-warmed ethanolamine blocking solution, cover with foil, and incubate for 30 min with slow and gentle shaking at RT.
   4. Discard the ethanolamine blocking solution into the appropriate biohazard trash receptacle and place the slide in a new, clean staining tube filled with 50 °C DIW. Cover with foil and incubate for 10 min with slow and gentle shaking at RT.
   5. Discard the water and transfer the slides to a glass staining holder (Supplementary Figure 8B). Place the glass staining holder into a swinging plate holder and centrifuge dry the slide for 5 min at 200 x g and RT.
4. Set up a divider and proceed with the biological blocking.
   1. Place the slide onto a 16-well divider (Supplementary Figure 8C).
   2. Prepare PBST washing solution: 50 mL of phosphate-buffered saline (PBS), pH 7.4 + 0.1% Tween-20.
   3. Prepare biological blocking solution (PBS-OVA): 5 mL of PBS, pH 7.4 + 1% chicken ovalbumin (10 mg/mL).
      NOTE: The choice of blocking reagent is critical for the success of the detection of anti-Neu5Gc antibodies. Ovalbumin is made in chickens which, like humans, cannot synthesize Neu5Gc, causing a lack of expression of this Sia. For the detection of Neu5Gc, even minor contaminants can dramatically reduce the sensitivity of the assay, sometimes even failing to detect any anti-Neu5Gc reactivity. For example, the most commonly used blocking reagent, bovine serum albumin (BSA), although not glycosylated by itself, contains bovine IgG contaminants that carry Neu5Gc-moieties and therefore compete with printed glycans when binding to sample with anti-Neu5Gc antibodies^19,20.
   4. Prewet the wells. Using a multi-pipette, aliquot 200 µL of PBST into the slide-wells, place them in a covered humid chamber, and shake for 5 min.
      NOTE: The humidity chamber is a glass tray with a wet towel paper, covered with nylon wrap and foil to protect the samples from light.
   5. Remove the PBST by flicking and tapping gently on a clean paper towel and aliquot 200 µL of PBS-OVA blocking buffer into each well. Place in a humid chamber and incubate for 1 h at RT with gentle shaking.
5. Prepare primary detection diluted in PBS-OVA blocking buffer, as listed in Table 2.
   NOTE: For quality control (QC) assessment of this sialoglycan microarray, use several Sia binding proteins: Sambucus nigra lectin (SNA), isolated from elderberry bark, is expected to recognize Siaα26; Maackia Amurensis Lectin II (MALII) is expected to recognize Siaα23; and chicken anti-Neu5Gc IgY is expected recognize all Neu5Gc-containing sialoglycans. In addition, use various human sera samples to profile anti-Neu5Gc IgG response. The concentration of lectins/antibodies/sera should be calibrated experimentally.
6. Flick dry PBS-OVA blocking buffer and add 200 µL of primary antibody per well (minimum volume per well: 70 µL), incubate in a humid chamber for 2 h.
7. Flick dry the primary detection and wash 4 times with PBST (between the 3^rd and 4^th wash, incubate the slide in PBST for 5 min in the humid chamber). Wash once with PBS.
8. Prepare the secondary antibody in PBS, as listed in Table 2.
9. Flick dry the PBS and add 200 µL of secondary antibody. Incubate for 1 h at RT with shaking.
10. Flick dry the secondary antibody and wash four times with PBST (between the 3^rd and 4^th wash, incubate the slide in PBST for 5 min in the humid chamber).
11. Carefully remove the frame and place the slides into a 50 mL slide staining tube filled with PBS and shake for 5 min.
12. Fill two slide-staining baths (Supplementary Figure 8D) with DIW, place the slide into the slide-holder, and dip it inside the bath. Quickly dip 10 times and then transfer to the second bath and dip 10 more times. Incubate for 10 min at RT with shaking.
13. Discard the water and transfer the slide to a glass staining holder (Supplementary Figure 8B; allow the slide to air dry). Place the glass staining holder in a swinging plate holder and centrifuge dry the slide for 5 min at 200 g and RT.
14. Scan the slide and then store it in a vacuum-sealed box in the dark.

6. Array Scanning and Data Analysis

1. Open the fluorescent slide scanner and let it warm for 5 min. Open the scanning and analyzing software.
2. Set the scanning parameters (Supplementary Figure 9A): scan at 532 nm and 100% power with a gain of 350 PMT, with one line to average and a 0-µm focus position. Use a 10.0 µm pixel size resolution.
   NOTE: Scanning parameters can be adjusted to different slide types, fluorescence, and resolution.
3. Scan the developed slide. Open the scanner door and place the slide inside, array facing down. Start scanning (green "Play" button on the software navigation panel menu, Supplementary Figure 9B).
4. Save the scanned slides as a tiff file.
5. Prepare slide for analysis. Select "Load array list" and upload the appropriate GAL file that maps the arrays in each block on the slide (Supplementary Figure 9C).
   NOTE: The GAL file contains information regarding both the identity and location (X, Y) of each spot on the slide. This file can be created by the printer software exporting a tab-delimited text file based on samples identities in the 384-well source plate and the defined print design.
6. Set the alignment and analysis parameters. Press the "Option" button on the software navigation panel menu (Supplementary Figure 10).
7. Define the alignment parameters. Find circular features, resize features during alignment by 50 – 350%, limit feature movement to a maximum translation of 40 µm, unflag features that fail background threshold (CPI threshold 0), estimate the wrapping and rotation when finding blocks, and automate image registration (10 pixels) (Supplementary Figure 10).
8. Define the analysis parameters: W1/ 532 ratio, normal standard deviation, analyze absent features, and background subtraction. Click "select" and then "Local feature background median" and set 2 pixels to exclude and a 3-pixel width of background. Press "OK." Set the scanner saturation level to default (Supplementary Figure 10).
   NOTE: The background subtraction method depends on the experimental design and can be modified according to specific needs.
9. Align the features maps. Set the program into block mode (press on "B"), select all blocks (Ctrl+A), position the array map grids, and align all blocks (Alt+Shift+F7) (Supplementary Figure 11).
10. Refine the feature alignment in each block. Zoom (press "Z" to get into zoom mode) into the upper-left block, press "B" again (block mode), and press the grid of this block. Only move the grid of this block until it aligns perfectly with the block spots and press F5 to align the features in this selected block. Repeat the grid movement and F5 alignment until the spots are perfectly circled. Do the same for every one of the 16 blocks until all grids are in place.
11. Analyze and save the data. Select all blocks (Crtl+A) and then analyze the data (Alt+A). Save the analysis results (Ctrl +U) as a .gpr file.
12. Open the saved .gpr file in a spreadsheet program and analyze the intensity of each spot based on fluorescence after local background subtraction (F532-B532).