Following is a written protocol for SEPCON assembly.  (Here is a link to a video demonstration)

Materials required: wafer tweezers, small sample tweezers (preferably plastic), soft foam board from craft store, wafer of sepcon chips, razor blade, plastic enclosure to store complete SEPCON, top basket, bottom basket,  350 µm thick silicone sheet of pre-cut “U” and square shaped gaskets.  (outside length – 5.5 mm, inside length – 4.0 mm)

SEPCON assembly supplies

From left to right: Wafer tweezers, metal chip tweezers, plastic sample tweezers

Popping chips out of a wafer is a time consuming and delicate task.  The membranes are relatively fragile and the forces applied to break out the chips can lead to their destruction.  Begin by slightly elevating the wafer on one side using the wafer tweezers.  (a few millimeters may suffice)

Next,  glide the plastic tweezers along the seams between chips while gently applying pressure.  Try to identify a weak seam and focus on that region, removing a section comprising multiple chips as opposed to removing a single chip.  It may take several passes, gradually increasing pressure as the seams are weakened.  The greater the pressure applied, the more likely an abrupt separation will take place which often results in membrane damage and debris creation.

Plastic tweezers popping out section of chips from slightly elevated wafer

Once a section with multiple chips has been separated from the full wafer, place the wafer back in its holder and focus on the section of chips.  Using the wafer tweezers, elevate one side of the section and again apply pressure with the plastic tweezers at the seams between chips.

Single SEPCON chip after being separated from small subsection of wafer.

Now that the chips are separated, it is necessary to visually inspect the membranes using an optical microscope.  It may be possible to tell by eye if there is damage, but very small defects will only be visible under the microscope.  Below are example images showing a perfect 5 slot SEPCON  and a damaged SEPCON.  The slot with a  broken membrane is allowing the front side illumination to pass directly through, making the slot look black.

Perfect SEPCON

Damaged SEPCON

Below are backside illuminated images which are useful sometimes, depending on the membrane being inspected.  (The camera reduced the quality of the image significantly)  In this case, it is nearly impossible to differentiate between the intact membranes and the broken membranes.

The gaskets below are cut from a sheet of silicone ???? µm thick using a Silhouette brand cutter.

Silicone sheet of gaskets

Place a “U”-shaped gasket into the bottom basket oriented with the open ends facing the same direction.  This will allow filtrate to flow out of the the assembly once completed.

Once the “U”-shaped gasket is firmly and evenly placed in the base of the bottom gasket, place a SEPCON chip on to with the “bottom” side up and the slots running towards the opening in the basket.  The “bottom” of the SEPCON can be determined by looking for the indented, trench-like look of the slots.

Next, place the square gasket on top of the SEPCON chip, trying to keep the gasket flat and centered.

chip into bottom basket on top of u-shaped gasket

Next, the top basket is snapped into the bottom basket.

Note: The square base of the top basket has three sides with longer plastic ridges that will lock into the bottom gasket.  One side has a shorter ridge and this must be orientated towards the open end of the bottom basket to allow for fluid to escape the assembly.

Try to put evenly distributed pressure on the top basket as it snaps down into the bottom basket.

top basket snapped into bottom basket

Next, cut an “X” in the cap of the  plastic housing.  (This is to ensure that a pressure differential isn’t created between the top and the bottom of the SEPCON.)

Finally, place the SEPCON assembly into the plastic housing and seal with the cap.

final assembled device

SEPCON assembly

I got some experiences in dealing with cells. This is for your reference.

 Aseptic Technique and Good Cell Culture Practice 

1. Sanitize the cabinet using 70% ethanol before commencing work.
2.  Sanitize gloves by washing them in 70% ethanol
3.  Put all materials and equipment into the cabinet prior to starting work after sanitizing the exterior surfaces with 70% ethanol.
4.  While working, do not contaminate hands or gloves by touching anything outside the cabinet (especially face and hair). If gloves become contaminated re-sanitize with 70% ethanol.
5. Equipment in the cabinet or that which will be taken into the cabinet during cell culture procedures (media bottles, pipette tip boxes, pipette aids) should be wiped with tissue soaked with 70% ethanol prior to use.
6. Movement within and immediately outside the cabinet must not be rapid. Slow movement will allow the air within the cabinet to circulate properly.
7. After completing work disinfect all equipment and material before removing from the cabinet. Spray the work surfaces inside the cabinet with 70% ethanol and wipe dry with tissue.
8. Sanitize the cabinet with UV light.

Make the media (10% FBS media) 

1. Autoclave a 250ml flask
2. The complete media consists of:

•                  250ml 1X DMEM (Cellgro: 10-013-CV)
•                  25ml FBS (Gibco: 26140-087)
•                  1ml L-Glutamine ( Gibco: 25030-081)
•                  1ml penicillin streptomycin (Gibco: 15140-122)

Use a Nalgene rapid-flow sterile disposable filter to put those contents in the flask. 

Thawing Cells

1. Pipette 9 mL of warmed complete media into the T75 flask.
2. Take the cryogenic vial from the liquid nitrogen and place in the 37℃ water bath to thaw.  Keep the cap above the water level, and move the vial through the water to aid in the thawing process.  Thawing should take less than a minute.
3. Pipette contents of the vial into the flask.
4. Rock the flask back and forth to distribute the cells.
5. Place the flask in the incubator.
6. Label T75 flask with cell type, passage number (this could be #0), date, and group member’s initials.

Changing Media 

1. Put the warmed complete media in the hood.
2. Aspirate the media in the flask using the glass Pasteur pipette.  Place the pipette a little away from the growth surface, to reduce the chance of sucking up cells.
3. Pipette 10 mL warmed complete media into the flask.
4. Close the lid and put the flask back in the incubator.

Passaging Cells (1:4 split)

Cell Culture Splits – a 1:4 indicates that one fourth of the cells are transferred into a new flask. There is no magic number or formula for how to do this.

1. Pipette 7.5 mL of warmed complete media into the T75 flask. Incubate the flask in the incubator for a while to achieve good CO2 content and pH.
2. View cultures using an inverted microscope to assess the degree of confluency and confirm the absence of bacterial and fungal contaminants.
3. Aspirate the media in the flask using the glass Pasteur pipette.
4. (Omitted) Wash the cell monolayer with 1-2 ml of PBS without Ca2+/Mg2+ (CMF-PBS).
5. Pipette trypsin onto the cells (eg. 1 ml for T25 flask, 2 ml for T75 flask). Rotate flask to cover the monolayer with trypsin and incubate for 3-4 minutes.  Check your flask and if the cells are not detached, incubate for another 2 minutes.
   • 0.25% Trypsin (Gibco: 15050-065)
   • Detached cells will have a rounded morphology and will float in the trypsin.
   • You can gently tap the flask to encourage the detachment process.
   • Cells should only be exposed to trypsin long enough to detach cells. Prolonged exposure could damage surface receptors
   • You may examine the cells using a microscope to ensure that many (~40%) the cells are detached and floating.
6. Pipette 8ml fresh medium to inactivate the trypsin. Pipette the solution up and down several times. (CAREFUL not to aspirate your media into the pipet aid)
7. Check the flask to ensure very few cells are left behind.  If there are remaining cells, wash the surface again with media.
8. Transfer the required number of cells (2.5ml for 1:4 split) to a new flask containing pre-warmed medium.
9. Pipette enough media to have a total of 10 mL of volume in your flask.
10. Rock the flask back and forth to distribute the cells in the flask, and then place in the incubator.
11. Label the flask with the cell and media type, passage number, cell split rate, date, and your initials.

Seeding cells on CytoVu 

1. Pre-condition the membrane by applying media (10% FBS), 25 μl to the basal wells and 10 μl to apical wells.
2. Replace lid and incubate at least 2 hours.
3. The same procedure with cell passaging (step 1-7). Suspended cells could be pipette to 15 mL conical vial.
4. Aspirate media using hand-held pipettes.
5. Add 10 μl of suspended cells to apical well.
6. Add 25 μl of media in basal well.
7. Replace lid and allow cells to attach to membrane.
8. Incubate to let cells grow to reach confluence. Change media every day. 

Results:

These are the pictures of cells on cytovu. Cells in 3 wells were alive. At first, the cells seemed to be very unhappy with the membrane. But things were better after day2 (I changed media every day).

Fabricate ITO Electrodes

1. Deposit 2500 Angstroms, ITO, Room Temperature Using Kurt Lesker Sputterer (Use ITO Variable Recipe, takes about 1.5hr)
   1. Tape in Microscope Slides and Coverglass into Machined Shadow Mask using Kapton Tape
2. Anneal in Toaster Oven, 450 Fahrenheit, 1hr
   1. Drops resistance from > 40 Megaohm to < 5 kohm. Measure from edge of Pad to Middle Finger

   2. 

      Hotplate Anneal Attempt of ITO Electrode (failed).

3. Clean Slides with Alcohol
4. Measure Resistance and record on side of slide. Check for Shorts across the pads
   1. 

RTP pnc-Si filters

1. Place susceptor and lid gently into RTP tool
2. Run warm up recipe on the RTP
3. Wait for the Oven to cool below 300 C
4. Remove susceptor and lid, take off lid
5. Place samples in susceptor, flat side up (blue), cover with lid
6. Place susceptor and lid gently into the RTP tool
7. Use the Simpore Post anneal recipe 
8. Wait for the Oven to cool below 300 C
9. Remove Susceptor, take off lid
10. Remove samples, place lid back on susceptor
11. Wait for the oven to cool down to room temperature, then turn off tool

Design Gaskets

1. Use adobe illustrator to draw your design, with the thinnest line width possible.
2. When you are satisfied, save an adobe illustrator file (.ai)
3. Once saved, use the export command to create a (.dxf) file. Use the default settings (make sure maximum editability is checked, and the 1 mm = 1 unit is accurate)
4. Email the .dxf file to yourself, or sneakernet it downstairs to the windows pc of the McGrath Computational alcove
5. Open Robomaster
6. Drag and drop the .dxf file onto the Robomaster window. Open-> file will not work.
7. The design will be offset, highlight all the components of your design and drag the design into the active area of the Robomaster window
8. Export as a .GSD file

Fabricate Gaskets

1. Use Silhouette Cutter in McGrath Computational alcove, preloaded with designed GSD File.
2. Wear gloves to keep things more cleanly
3. Place silicon sheet with coverplastic attached onto cutting mat.
4. Align and load cutting mat into cutter by pressing “load cutting mat”
   1. If you choose “load media” your cuts will not line up with the grid in software.
5. When you have positioned your design and are ready to cut, select “send to silhouette”
   1. Check the Custom Settings. For Silicone sheets with coverplastic (custom media), the general settings are Blade:5, Speed:25, Thickness:5, using cutting mat and double-cut
   2. Double check the height of the blade by removing it from the holder. If the height is wrong, adjust it using the ring on the base of the cutter, then reinstall the blade
   3. If you are unsure of your cut settings, a test cut can be performed. Use the software arrow keys to position the blade over a uncut area of silicone and press test cut. The test is successful if you can pick up the triangle without removing the square.
6. Press Cut
7. Eject the cutting mat by pressing enter
8. Remove the cover plastic from the cut areas, then lift out your cut gaskets.
9. Replace the coversheet on top of the cutting mat.

Device Assembly

1. Prepare a 10:1 PDMS to Curing Agent mixture (2g is plenty)
   1.  Outgas for 20 minutes in vacuum can
2. Mortar bond Channel gaskets to Electrodes
   1. Take Top gasket and mortar bond by gently rubbing a light coat of PDMS using green applicator onto bonding surface, then join to top electrode (slide with holes), where the electrode surface and the PDMS coated surface are joined. Line up Outer holes with top gasket ports. Gently massage out air bubbles.
   2. Take Bottom gasket and mortar bond by gently rubbing a light coat of PDMS using green applicator onto bonding surface, then join to bottom electrode (thin coverglass) where the electrode surface and the PDMS coated surface are joined. Center gasket onto electrode pattern. Gently massage out air bubbles.
   3. If we can get access to a spin coater, the manual application could be refined to a “stamp and stick method”
3. Outgas both substrates for 5 minutes in vacuum can
4. Cure PDMS with hotplate, 150 C for 10 minutes. Use glass cover over samples
5. Mortar bond Interstitial Gasket
   1. Take Interstitial gasket and mortar bond by gently rubbing a light coat of PDMS using green applicator onto bonding surface, then join to top gasket stack, where the uncoated top gasket and  the PDMS coated interstitial gasket surface are joined. Line up ports with top gasket ports. Gently massage out air bubbles.
6. Mortar bond Filter
   1. Mortar bond by gently rubbing a light coat of PDMS using green applicator onto exposed top surface gasket, taking care not to get PDMS into the channel,  then join to filter (blue side down, green side up), where the uncoated blue filter surface and  the PDMS coated top gasket surface are joined. Center filter in area, making sure top ports are occluded and the filter channel lines up with the gasket channel. Press down gently.
7. Outgas in vacuum can for 5-10 minutes (more because harder to massage out air bubbles)
8. Cure PDMS with hotplate, 150 C for 10 minutes. Use glass cover over samples.
9. Fill in Filter/Interstitial gasket gap with PDMS using thin pipette tip
   1. Take a small pipette tip and get a small drop of pdms on it. Rub all edges of the filter, returning to get a new drop periodically, filling up the slot. Some may spill out over the sides; be as cleanly as possible. Spread out run off onto gasket, not the filter .
10. Outgas in vacuum can for 5-10 minutes (more because harder to massage out air bubbles)
11. Cure PDMS with hotplate, 150 C for 10 minutes. Use glass cover over samples.

    1. 

       Filter and interstitial cure

12. Mortar bond stack to bottom gasket/electrode.
13. NO OUTGASSING!!! WILL DRAW PDMS INTO CHANNEL
14. Cure PDMS with hotplate, 150 C for 10 minutes. Use glass cover over samples.
15. Clean exposed glass surface with alcohol wipe.
16. Use ozone bonding (15 minutes) to join PDMS feet (pre drilled holes) to microscope slide.
17. Bake on hotplate 150C for 10  minutes to finish fabrication