Surface Chemistry Protocol


PROTOCOL POPA LAB (last update June 2021)


  • (3-aminopropyl)trimethoxysilane (referred to as Silane, stored in desiccator), 97%, Sigma-Aldrich, Cat. No. 281778-5ML
  • Acetone HPLC, EMD Millipore, Cat. No. AX0115
  • Amino Polystyrene Particles, 5% w/v, Spherotech
  • 5-2.9 µm, Cat. No. AP-25-10
  • 0-3.4 µm, Cat. No. AP-30-10
  • 5-3.9 µm, Cat. No. AP-60-10
  • Casein blocking buffer, 3% in PBS, Fisher Scientific, Cat. No. AAJ60289AK
  • Glutaraldehyde, 70% aqueous (stored at -20 0C), Sigma-Aldrich, Cat. No. G7776-10mL (!CAUTION: Hazardous chemical. When handling, wear gloves and safety glasses. Work under hood and do not inhale vapors)
  • HaloTag Amine O4 ligand, 5mg/mL dissolved in DMSO (stored in small aliquots at -80 0C under Argon), Promega (Cat. No. P6741) & Sigma-Aldrich (Cat. No. D2650-5X5ML)
  • Hellmanex (TM) III, Sigma-Aldrich, Cat. No. Z805939-1
  • Methanol (MeOH), Fisher Scientific, Cat. No. A413-4
  • Micro Cover Glasses, 22 x 22 x (0.13 – 0.16) mm thick, Ted Pella, Cat. No. 260140
  • Painter’s Touch® 2x Ultra Cover® Spray Paint, Rust-Oleum, Cat. No. 249127
  • PLA+ 3D Printer Filament 1.75mm, SunLu PLA Filament Pro,
  • Potassium Chloride Anhydrous (KCl), Sigma-Aldrich, Cat. No. 793590-2.5KG
  • Sodium Phosphate Dibasic, Na2HPO4, Sigma-Aldrich, Cat. No. S5136-500G
  • Sodium Phosphate Monobasic, NaH2PO4, Sigma-Aldrich, Cat. No. RPI S23185-500.0
  • Superslip® Cover Glasses, 24 x 40mm x 0.13 – 0.16mm thick, Ted Pella Cat. No. 260164
  • Sulfhydryl blocked-BSA, Lee Biosolutions, Cat. No. 100-10SB
  • Superparamagnetic Dynabeads®M-270 Beads, Thermo Fisher Scientific
  • Streptavidin coated, No. 65305
  • Amino groups, No. 14-307-D
  • Sylgard™ 184 Silicone Elastomer, Dow Corning, Cat. No. 4019862

NOTE: All aqua solutions are made with double-distilled water.


  • Analog Lab Oven, Quincy Lab, No. G4250242
  • Glass Berzelius Beakers 250 mL (x2)
  • Columbia Staining Jars, DWK Life Sciences (Wheaton), Cat. No. W900180
  • Coplin Staining Jars, 304.8 x 228.6 x 152.4 mm, DWK Life Sciences (Wheaton), Cat. No. 900570
  • Corning® 5 x 7 Inch Top PC-400D Hot Plate with Digital Display, Cat. No. 6795-400D
  • Plasma Cleaner, Harrick Plasma, No. PDC-32G
  • Prusa I3 MK3S+ 3D printer, Prusa Research
  • Tube Rotator, Argos Technologies® RotoFlex, Cole-Parmer, Cat. No. EW-04397-33
  • Staining Rack, Thomas Scientific, Cat. No. 24957
  • Ultrasonic Cleaner, Branson Ultrasonics, No. 2210R-MT


  • PBS buffer: 50 mM Na2HPO4 / NaH2PO4 and 150 mM KCl (25 ml of 0.5 M Na2HPO4 stock sol, and 12.5 ml of 1M NaH2PO4) (pH 7.2)
  • TRIS/BSA buffer: 20 mM Tris-HCl, 150 mM KCl, and 1% w/v Sulfhydryl blocked-BSA (pH 7.4)




  1. Place nine 24X40 mm coverslips (surfaces) in each of the two Coplin jars.
  2. Sonicate the surfaces for 20 min in 1-2 % aqueous Hellmanex III solution at 50-60 o
  3. Rinse (at least) 10-15 times with ddH2O.

CRITICAL STEP: Ensure that ALL detergent is washed and the water does not make any bubbles.

  1. Sonicate the surfaces in acetone for 20 min at RT with the acetone level just above the upper edges of the surfaces.
  2. Rinse once with methanol.
  3. Sonicate the surfaces for 20 min at RT in methanol
  4. Air-dry the surfaces and move them from the staining jar to the staining rack, facing opposing orientations. Further dry the surfaces in the oven at 110 oC for ~10 min. Simultaneously, rinse thoroughly with methanol the two glass Berzelius beakers thoroughly with methanol (one is used for silanization, second for washing) and dry them face-down in the oven at 110 oC.


  1. Place the staining rack with the dried surfaces in the plasma generator for 20 min to activate them.

NOTE A: Caution, the staining rack might be hot. Use tweezers. Also always use gloves when handling the staining rack.

NOTE B: After turning on the vacuum pump connected to the plasma cleaner, open slowly the valve to let some air into the chamber until the plasma color is at its brightest intensity (purple color for air plasma) to ensure proper activation.

  1. While the surfaces are being activated in the plasma cleaner, prepare 0.1% solution v/v of silane in methanol (150 mL silane in 150 mL methanol).

NOTE: Make solution in a 50 mL tube and mix it well before adding it to the glass beaker/silanization chamber. Bring the beaker close to the plasma cleaner. 

  1. Following the 20 min plasma-activation step, release the chamber vacuum and quickly move the rack with tweezers from the plasma cleaner chamber into the beaker with silane/methanol. Leave in the silanization beaker for 20 min.

CRITICAL STEP: Move the rack with coverslips from the plasma cleaner chamber into the silane solution as fast as possible, desirably in less than 10 s, as the glass deactivates quickly in air.

NOTE: Use fresh silane (don’t purchase volumes larger than 5 mL). Typically a silane stops working properly after ~1 month, when kept in the desiccator. Toward the end of its 1-month life, the silanization time can be decreased to 10 min if beads stick non-specifically during experiments.

  1. Move the rack in the second wash-beaker, which contains only methanol and slowly shake it to wash away non-reacted silane. Then leave the rack in the washing beaker for 2 min.
  2. Air-dry the silanized surfaces with an air gun and place them in the oven at 110 oC for at least an hour.
  3. After 1+ hours. either move the silanized coverslips to a desiccator for storage (up to two weeks) or proceed to the fluid-chamber assembly step.



  1. Place seven 22X22 mm coverslips in each of the three Columbia staining jars.
  2. Sonicate for 20 min in 1-2% aqueous Hellmanex III solution.
  3. Rinse 10-15 times with ddH2O until water runs clear.
  4. Rinse with methanol once.
  5. Dry with air and place the jars (without their plastic lids) inside the oven at 110 oC for 15-20 minutes, for further drying.


  1. Arrange the coverslips in rows on an A4 paper that has 22X22 mm squared boxes printed on it.
  2. Fix the coverslips on that paper with labeling or masking tape, covering about 3 mm of the top and bottom edges. Place the paper inside a cardboard box.
  3. Gently spray black Rust-Oleum paint over the coverslips in the fume hood maintaining a ~40 cm (~15 in) distance between the spray can and surfaces.

! CAUTION: Spray aerosols can be toxic or can stain your clothes. Do this step under fume hood.

  1. Leave surfaces to dry under fume hood for 15 min and spray another coat. Leave to dry for another 30 min.


  1. Mount six surfaces at a time on the 3D printer plate following the grid (starting from SW corner) with paper tape.
  2. Select the “6chamberMT_cover_100microns” program from the 3D printer menu and wait for the printer to complete depositing the PLA spacing material.

NOTE: During printing, if coverage is not homogeneous, try to decrease speed or change temperature or extrusion percentage from the printer menu.

  1. Set the hot plate to 250 o Place a cover slip ontop of the silanized bottom glass with printed chamber and put the two on the hot plate. Gently press with the back of a metal blade to seal the chamber.
  2. Store the assembled fluid chamber in the desiccator (for up to two weeks since silanization was done) or proceed to the next step.

SURFACE FUNCTIONALIZATION (TIMING ~ 5 hours/overnight + 12 hours):

  1. Prepare 1% Glutaraldehyde solution v/v in PBS (20 mL of 70% Glutaraldehyde in 1.3 ml PBS). Vortex the amine-terminated polystyrene beads solution (reference beads for single molecule experiments) of desired size and add 6 m Then vortex the Glutaraldehyde-reference beads mix.

NOTE: Glutaraldehyde solution is viscous, so you need to cut the end of a 200 mL tip and only slightly immerse it in the stock solution, so residue does not form on the outside wall of the tip.

! CAUTION: Handle glutaraldehyde under the fume hood, as it is toxic. Wear glass and protective equipment.

  1. Place fluid chambers on a folded napkin under a 30-45 deg angle and add a total of 100-200 mL at the top end of the chamber (typically you need to add ~50 mL and wait for the solution to flow before adding the rest).

NOTE: Make sure the solution flows through the fluid chamber, and not over it. You can use a napkin at the bottom end to aid it to flow, or a pipette tip at the top end, to generate small flow.

  1. Then straighten the chambers and add either PBS buffer or the remaining glutaraldehyde-reference beads solution at both ends of the chamber, to prevent drying. Leave to react for 1 hour.
  2. Tilt again the fluid chamber and was with 200 mL of PBS to remove, the unreacted glutaraldehyde and unbound polystyrene beads.
  3. Prepare 10 mg/mL HaloTag O4 solution (500x dilution: 1.3 ml PBS + 2.2 mL ligand 5 mg/mL) and add 100-200 mL in each chamber for 4 h at RT or overnight at 4 o
  4. Then straighten the chambers and add either PBS buffer or the remaining HaloTag O4 solution at the ends of the chamber. Move the chambers from the fume hood, to prevent evaporation.
  5. Quench the reaction by passing through the chamber 200 mL of TRIS buffer.
  6. Passivate the chambers with 1% BSA in TRIS/KCl solution for 12 h before use.


  1. Wash passivated chamber with 200 mL of PBS.
  2. Freshly dilute thawed protein of interest from -80 oC to a concentration of 250-300 nM in PBS.

HaloTag – SpyTag terminated proteins

  1. Incubate the chamber with HaloTag-SpyCatcher protein 10 mM diluted in PBS, for 10 minutes at RT. Then wash with ~1 mL of PBS.
  2. Incubate the chamber with 50 mL of the diluted protein for 30-45 min at RT. Then wash with ~1 mL of PBS.
  3. Vortex paramagnetic beads functionalized with HaloTag and add 20 mL in the fluid chamber. Then place chamber ontop of microscope and add a drop of PBS buffer at each end, to prevent drying.

HaloTag – AviTag terminated proteins

  1. Incubate the chamber with 50 mL of the diluted protein for 10-15 min at RT. Then wash with 200 mL of PBS.
  2. Vortex Streptavidin-coated paramagnetic beads functionalized and add 20 mL in the fluid chamber. Then place chamber ontop of microscope and add a drop of PBS buffer at each end, to prevent drying. 


STREPTAVIDIN-COATED BEADS (TIMING 20 min + overnight + 20 min):

  1. Add 10 mL of Streptavidin coated paramagnetic beads to 200 mL PBS in a 1.5 mL tube (or 20 mL of Streptavidin with twice the buffer or solution at every step).
  2. Vortex the bead solution and wash 3x with the same volume of PBS by sedimenting the beads with a magnet, gently removing the solution while holding the magnet close to the 1.5 mL tube.

NOTE: Make sure that the beads are not being sucked during the buffer removal step.

  1. Block the paramagnetic beads with 200 mL of the 1-1.5% Casein solution overnight at 4 oC in the rotating platform.
  2. Next day, wash the beads again with PBS for 3x and leave them on the rotating platform until you are ready to use them.

HALOTAG LIGAND-COATED BEADS (TIMING 1.5 hours + overnight + 24 hours):

  1. Add 20 mL amine-terminated paramagnetic beads to 650 mL PBS in a 1.5 mL tube and vortex.
  2. Wash the beads 3X with PBS (650 mL) buffer sedimenting the paramagnetic beads from the solution using a strong permanent magnet.
  3. Suspend the beads in 1% Glutaraldehyde solution v/v (10 mL of 70% Glutaraldehyde in 650 ml PBS) and rotate for an hour at RT.
  4. Wash the beads 3X with PBS buffer (650 mL).
  5. Dilute HaloTag O4 ligand in PBS (1.1 ml of 5 mg/ml ligand + 650 ml PBS). Remove the PBS buffer from tube while keeping the beads sedimented with a magnet and add the HaloTag O4 solution. Vortex tube and leave beads to react with the HaloTag ligand overnight, while spinning in the tube rotor at 4 oC.
  6. Next day wash the beads with PBS and passivate for 24 hours with 200 mL of the Casein solution 1-1.5% w/v, while spinning in the tube rotor at 4 oC.
  7. Wash the beads 3x with PBS and leave them on the rotating platform until you are ready to use them.



This protocol is inspired from:
Nat. Protocols, 2013, 8 (7): 1261-1276, link
J. Am. Chem. Soc., 2016, 138 (33), 10546–10553, link
Phys. Chem. B, 2020,  124 (16), 3283-3290, link.