Hanging Cell Culture Inserts

Note: Depending on the cell lines and the nature of the co-culture, the researcher can decide which side of the membrane is seeded first. To maintain sterile incubations of cells seeded on the underside of the membranes, use sterile Petri dishes for the Millicell® single-well inserts and sterile feeder trays for Millicell® 24- and 96-well cell culture insert plates.

1.   Using an optimized seeding density, seed the first cell type in either the apical wells or on the basolateral underside of the membranes. Refer to the recommended working volumes chart (see page 42) for appropriate apical volumes. For basolateral seeding volumes, use approximately 200 ìL for Millicell® 12 mm single-well inserts and Millicell®-24 insert plates, and approximately 30 µL for Millicell®-96 insert plates.
2.    Incubate in a 37°C CO2 incubator for 1 to 4 hours to allow the cells to attach to the membrane.
3.    Gently turn the Millicell® device over and seed the second cell type in either the apical well or on the basolateral underside of the membrane.
4.    Incubate in a 37°C CO2 in the incubator for 1 to 4 hours to allow the second cell type to attach.
5.    Add appropriate volume of media to the apical or basolateral chambers and return to incubator.

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A. Day 1
1.   Coat T-75 flasks with 10 mL of 0.1% gelatin in DPBS and incubate for at least 30 minutes at 37°C.
2.   Thaw PMEF vial (s) quickly in a 37°C water bath, transfer to 15 mL tube already containing 10 mL warm medium. Gently invert tube, and pellet cells at 4°C, ~1000 rpm for approximately 4–5 minutes.
3.   Remove supernatant, resuspend cells, remove gelatin from plates/flasks, and aliquote feeder cell suspension per densities recommended below.
4.   Remove excess gelatin from flask prior to seeding.
5.   Seed flask with MEF feeder cell suspension: approximately 1.5 × 105 cells per mL MEF should result in 95% confluence within 24 hours.
6.   Incubate at 37°C overnight.

B. Day 2
1.   Coat T-75 flasks with 10 mL of 0.1% gelatin in DPBS and incubate for at least 30 minutes at 37°C.
2.   Thaw PMEF vial (s) quickly in a 37°C water bath, transfer to 15 mL tube already containing 10 mL warm medium. Gently invert tube, and pellet cells at 4°C, ~1000 rpm for approximately 4–5 minutes.
3.   Remove supernatant, resuspend cells, remove gelatin from plates/flasks, and aliquote feeder cell suspension per densities recommended below.
4.   Remove excess gelatin from flask prior to seeding.
5.   Seed flask with MEF feeder cell suspension: approximately 1.5 × 105 cells per mL MEF should result in 95% confluence within 24 hours.
6.   Incubate at 37°C overnight.

C. Day 3
1.   Feed ESC on MEF feeder layer with fresh ESC media.

D. Day 4-8
1.   Feed ESC on MEF feeder layer with fresh ESC media or pass cells, at a 1:2 ratio, if required. (After thawing ESC, 2–3 passages are preferred before seeding onto a Millicell®-24 or Millicell® 96-well plate. Both cell types are lifted at once and passed on to a new T-75 containing inactivated MEF.) ESC colonies grown on Millicell®-96 cell culture insert 1.0 PET membrane, stained for alkaline phosphatase activity, after culturing via indirect co-culture with ESC in apical well, at a 200 cell per well seeding density, and MEF in the single-well feeder tray.

E. Day 9
1.   Feed ESC on MEF feeder layer with fresh ESC media.
2.   Coat Millicell®-24 or Millicell®-96 single-well feeder tray with approximately 5–10 mL of 25 µg/mL fibronectin in DPBS and incubate for 45 minutes at room temperature.
3.   Remove excess fibronectin from single-well tray.
4.   Thaw MEF using protocol from Day 1, section A.
5.   Seed fibronectin coated single-well tray with MEF cell suspension: approximately 1.67 × 106 MEF cells per single-well tray will result in 95% confluence within 24 hours.
6.   Cover with lid and incubate single-well tray at 37°C overnight.

F. Day 10
1.    Lift ESC and MEF feeder cells from T-75 flasks:
2.   Wash flasks 2X with 10 mL of pre-warmed DPBS (incubate 1–2 minutes per wash).
3.   Remove DPBS and add 3 mL TrypLE and incubate at room temp for 2–3 minutes.
4.   Monitor detachment of cells with an inverted microscope and add ESC media to inactivate TrypLE.
5.   Mix well and wash flask wall to remove all cells from flask.
6.   Separate ESC from MEF feeder cells:
7.   Transfer ESC/MEF cell suspension to a new T-75 flask and incubate at 37°C for 45 minutes.
8.   Remove non-adherent cells (ESC) and transfer to another new T-75 flask and incubate at 37°C for 45 minutes.
9.   Remove non-adherent cells (ESC) again and seed cell culture filter plate wells with ESC suspension.
10.   Seed apical wells of the cell culture plates with ESC. Seed approximately 200–500 cells per well in 100 µL ESC media for Millicell®-96 plates and approximately 1000–1500 cells per well in 400 µL ESC media for Millicell®-24 plates.
11.   Remove media from the MEF seeded single-well trays and replace with approximately 28–32 mL ESC media.
12.   Combine ESC seeded cell culture filter plates to MEF seeded single-well trays.
13.   Incubate assembly at 37°C overnight.

G. Day 12 and Day 14
1.   Feed ESC and MEF indirect co-culture with ESC media.

H. Day 16
1.   Analyze alkaline phosphatase activity to demonstrate that ESC is undifferentiated with an alkaline phosphatase detection kit.

Note: This protocol is designed to grow undifferentiated embryonic stem cells in an indirect co-culture with the fibroblast feeder layer. Although it is targeted for use with Millicell®-24 and Millicell®-96 plates, this protocol can be used with Millicell® single-well inserts as well.

Materials and Reagents

• Millicell®-24 cell culture insert plates — Millipore cat. nos. PSHT 010 R5, PSRP 010 R5
• Millicell®-96 cell culture insert plates — Millipore cat. no. MACA C02 B5
• Primary mouse embryo fibroblasts (DMEF) — Millipore cat. no. PMEF-CFL
• 129/S6 Murine embryonic stem cells (ESC) — Millipore cat. no. SCR012

ESC Media:

• Knock out DMEM — Millipore cat. no. SLM-220-B
• 20% ES qualified Serum — Millipore cat. no. ES-009-B
• 1% Glutamax-1
• 1% PenStrep — Millipore cat. no. TMS-ABZ-C
• 1% Non Essential Amino Acids — Millipore cat. no. TMS-001-C
• 0.1% ESGRO® (LIF) — Millipore cat. no. ESG1106
• 0.1% 2-mercaptoethanol — Millipore cat. no. ES-007-E

MEF Media:

• DMEM — Millipore cat. no. SLM-022-B
• 10% Fetal Bovine Serum — Millipore cat. no. ES-009-B
• 1% Glutamax-1
• 1% PenStrep — Millipore cat. no. TMS-ABZ-C
• 1% Non Essential Amino Acids — Millipore cat. no. TMS-001-C
• Gelatin 2% Solution — Millipore cat. no. SF008
• Mitomycin C powder — Sigma cat. no. M4287
• DPBS — Millipore cat. no. BSS-1005-B
• TrypLE™ Select (1X), liquid — Invitrogen cat. no. 12563

Tissue culture flasks and tubes

• Fibronectin Solution, 1mg/mL — Millipore cat. no. FC010

Other:

• Alkaline phosphatase detection kit — Millipore cat. no. SCR004

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Materials

• Millicell® inserts and plates.
• 70% Ethanol (filter sterilized with a Millex-GP filter unit . Millipore cat. no. SLGP 033 RS; or a Stericup-GP filter cup . Millipore cat. no. SCG-P U11 RE)
• Rat Tail Collagen, Type 1, approximately 3 mg/mL (Millipore cat. no. 08-115)
• Sterile pipette syringes and tips

Note: An alternate collagen source (Type I) of suitable concentration diluted in 0.01 N HCl or acetic acid may also be used.

Method
1.    Dilute the collagen 1:4 in 70% ethanol (1 part collagen and 3 parts 70% ethanol) and vortex until the collagen is solubilized.
2.    Gather the applicable number of Millicell® inserts or plates.
3.    Using a sterile pipette syringe, add the appropriate volume of the collagen/ethanol mixture to each well.
4.    Gently shake the cell culture plate until the collagen/ethanol mixture evenly coats the inside of the insert.
5.    Air dry inserts in a laminar flow hood. Leave cell culture plate cover ajar to allow airflow and prevent condensation. Note: Although drying typically takes anywhere from 3 hours to overnight, overnight drying is recommended.
6.    Seed the Millicell® insert with appropriate cell density (for example, MDCK cells: approximately 5×104 to 5×105 cells/cm2).

Volumes for Coating Millicell® Devices

Coating	(24-well)	(12-well)	(6-well)	(Millicell® 24)	(Millicell® 96)
Collagen (Collagen/Ethanol mix)	50 µL	150 µL	400 µL	100 µL	25 µL
Fibronectin* (Fibronectin/DMEM mix)	100 µL	300 µL	700 µL	200 µL	50 µL
Laminin* (Laminin/DMEM mix)	100 µL	300 µL	700 µL	200 µL	50 µL
Matrigel* (Matrigel/H2O solution)	100 µL	300 µL	700 µL	200 µL	50 µL

*Can be used on any Millicell® membrane
*Matrigel is a registered trademark of Becton, Dickinson and Company

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Materials

• Millicell® inserts and plates.
• Sterile pipette syringes and tips

Human Fibronectin, 1 mg/mL (Catalogue Number: 08-102) reconstituted. Handle and store according to manufacturer's instructions.

Method
1.    Dilute the fibronectin 1:10 in the serum-free DMEM (1 part fibronectin in 9 parts DMEM) and vortex until fibronectin is solubilized.
2.    Gather the applicable number of Millicell® inserts or plates.
3.    Using a sterile pipette syringe, add the appropriate volume of the fibronectin/DMEM mixture to each well.
4.    Gently shake the cell culture plate until the fibronectin/DMEM mixture evenly coats the Millicell®-CM insert(s).
5.    Air dry inserts overnight in a laminar flow hood. Leave cell culture plate cover ajar to allow airflow and prevent condensation. Note: Although drying typically takes anywhere from 3 hours to overnight drying is recommended
6.    Seed the Millicell® insert with appropriate cell density (for example, MDCK cells: approximately 5×104 to 5×105 cells/cm²).

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Materials

• Millicell® inserts and plates.
• Sterile pipette syringes and tips

Laminin, 1 mg/mL. Handle and store according to manufacturer's instructions.

Method
1.   Dilute the Laminin 1:10 in DMEM (1 part laminin in 9 parts DMEM) and vortex until the laminin is solubilized.
2.   Gather the applicable number of Millicell® inserts or plates.
3.   Using a sterile pipette syringe, add the appropriate volume of the laminin/DMEM mixture to each well.
4.   Gently shake the cell culture plate until the laminin/DMEM mixture evenly coats the Millicell® inserts.
5.   Air dry inserts overnight in a laminar flow hood. Leave the cell culture plate cover ajar to allow airflow and prevent condensation. Note: Although drying typically takes anywhere from 3 hours to overnight, overnight drying is recommended.
6.   Seed the Millicell® insert with appropriate cell density (for example, MDCK cells: approximately 5×104 to 5× 105 cells/cm²).

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Materials

• Millicell® inserts and plates.
• Sterile pipette syringes and tips

Method
1.   Dilute Matrigel according to manufacturer’s instructions.
2.   Gather the applicable number of Millicell® inserts or plates.
3.   Using a sterile pipette syringe, add the appropriate volume of the Matrigel mixture to each well.
4.   Gently shake the cell culture plate until the Matrigel solution evenly coats the Millicell® insert.
5.   Air dry inserts overnight in a laminar flow hood. Leave cell culture plate cover ajar to allow airflow and prevent condensation. Note: Although drying typically takes anywhere from 3 hours to overnight, overnight drying is recommended.
6.   Seed Millicell®-CM insert with appropriate cell density (for example, MDCK cells: approximately 5×104 to 5×105 cells/cm²).

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Consult the following table for chemical compatibility information with common fixative chemicals. 

	PS	HA	CM	PCF	PET
Acetic Acid	NR NR	NR NR	R R	R NR	NR Acetone R
Acetronite	NR	NR	R	NR	ND
Ammonium Hydroxide	TST R TST	NR NR NR	R R R	TST NR R	ND DMSO ND Alcohols R
Formaldehyde	NR	NR	R	R	R
Glutaraldehyde	RS R	ND R	R R	ND R	ND Glycerol R
Hydrochloric Acid, IN	R RS	R NR	R R	R R	R Methanol ND Sodium
Dodecyl Sulfate	ND	R	ND	TST	ND Sodium
Hydroxide, 3N	R	NR	R	NR	TST
TCA (aqueous solution)	ND	NR	R	TST	NR
Triton® x-100 Surfactant	R	R	R	R	R

PS = PS (polystyrene) Membrane 
HA = HA (mixed cellulose) Membrane
CM = CM (polytetra-fluoro-ethylene) Membrane
PCF = PCF (polycarbonate) membrane
PET = PET (polyterepthalate) membrane

General Considerations

• Compatibility key:
• R=recommended
• NR=not recommended
• RS=recommended for short term use
• TST=testing recommended
• ND=no data available
• If the chemicals are compatible with the membrane but not the polystyrene housing, remove the membrane from the housing before adding the chemical.
• Unless otherwise stated, the chemicals listed are at maximum concentration. If the plastic housing and/or membranes are not compatible with the maximum concentration, they might be compatible at lower concentration.

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Materials

• Millicell® inserts or insert plates
• Milli-Q® water
• Millex®-GP filter unit — Millipore cat. no. SLGP 033 RS
• Toluidine blue (Sigma)
• 3% glutaraldehyde in Phosphate Buffered Saline (PBS)
• Triton X-100 (Sigma), 0.5% Method

Method
1.   Prepare a 0.3 % solution (gram percent) of toluidine blue in Milli-Q® water, stir, and filter through a Millex-GP filter unit.
2.   Remove the Millicell® insert from the plate and wash gently with PBS to remove growth media.
3.   Fix the cells for 15 minutes with 3% glutaraldehyde in PBS.
4.   Rinse gently with Milli-Q® water. Repeat twice
5.   Permeabilize the cells with 0.5% Triton X-100 for 5 minutes.
6.   Rinse gently with Milli-Q® water. Repeat twice.
7.   Apply stain to the apical cell side of membrane for 30–60 seconds.
8.   Observe as a wet mount.

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The Hema-3® stain kit is a quick (less than 15 minutes) nuclear staining procedure that can be used with all Millicell® cell culture products. The kit is available through Fisher (cat. no. 22–122911).

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Materials

• Millicell®-CM cell culture plate inserts — Millipore cat. nos. PICM 012 50, PICM 030 50
• 3% glutaraldehyde in Phosphate Buffered Saline (PBS)
• Milli-Q® water
• 100% methanol
• Wright’s stain

Method
1.   Remove the Millicell®-CM insert from the plate and wash gently with PBS to remove growth media.
2.   Fix the cells for 15 minutes with 3% glutaraldehyde in PBS.
3.   Rinse gently with Milli-Q® water. Repeat twice
4.   Rinse once with methanol and incubate in fresh methanol for 5 minutes.
5.   Aspirate the methanol. Add Wright’s stain to cover the inside membrane.
6.   Incubate for 30 seconds.
7.   Rinse gently with Milli-Q® water. Repeat three times.
8.   Observe as a wet mount.

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Note: Standard histological hematoxylin and eosin staining techniques can be performed on thin sections.

Materials

• Millicell®-HA cell culture inserts — Millipore cat. no. PIHA 012 50, PIHA 030 50
• Millex®-GP filter unit — Millipore cat. no. SLGP 033 RS
• 3% glutaraldehyde in PBS, store at 4°C
• Phosphate Buffered Saline (PBS)
• 0.5% Triton X-100 (Sigma) in Milli-Q® water
• Hematoxylin solution, HHS-1, 7.5 g/L
• 0.5% Triton X-100 (Sigma) in Milli-Q® water
• Dilute ammonium hydroxide (8–10 drops concentrated ammonium hydroxide in 100 mL of water)
• 0.5% hydrochloric acid in 70% ethanol
• Milli-Q® water
• Cork borer

Method
1.   Filter hematoxylin solution through a Millex-GP unit and cover the cell layer.
2.   Incubate for 15 minutes at room temperature.
3.   Rinse the Millicell®-HA insert with Milli-Q® water to remove stain. Repeat twice.
4.   Destain by adding 0.5% hydrochloric acid in 70% ethanol for 2–3 minutes.
5.   Rinse with Milli-Q® water. Repeat twice.
6.   Add dilute ammonium hydroxide into the Millicell®-HA insert to cover the membrane. Incubate for 3 minutes or until a uniform blue color is observed on the membrane.
7.   Rinse with Milli-Q® water. Repeat twice.
8.   Using a cork borer remove membrane. Mount membrane on a slide in a commercial mounting medium. Make sure the cell layer is facing the microscope objective.

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Materials

• Millicell®-CM Culture Plate Inserts — Millipore cat. nos. PICM 012 50, PICM 030 50
• Millex-GP Filter Unit — Millipore cat. no. SLGP 033 RS
• Milli-Q® water
• Sterile Phosphate Buffered Saline (PBS)
• 3% glutaraldehyde in PBS, store at 4°C
• Methanol
• 0.5% Triton X-100 in Milli-Q® water
• Hematoxylin solution (Gill No. 1), filter sterilized with a Millex-GP unit (Millipore cat. no. SLGP 033 RS)
• 0.5% hydrochloric acid in 70% ethanol
• Dilute ammonium hydroxide (8–10 drops concentrated ammonium hydroxide in 100 mL of water)
• Mounting media

Method
1.   Remove the Millicell®-CM insert from the plate and wash gently with PBS to remove growth media.
2.   Add 3% glutaraldehyde (at 4°C) in PBS (pH 7.3) to the inside and outside (cell culture plate well) of the Millicell®-CM device. Leave for 15 minutes.
3.   Carefully remove glutaraldehyde. Add methanol to the inside and outside (cell culture plate well) of the Millicell®-CM unit. Leave for 10 minutes.
4.   Carefully remove methanol. Add 0.5% Triton X-100 to the inside and outside (cell culture plate well) of the Millicell®-CM unit. Leave for 5 minutes.
5.   Carefully remove Triton X-100. Add hematoxylin solution (Gill No. 1, Sigma Chemical, filtered with a Millex® unit). Leave for 15 minutes.
6.   Wash the Millicell®-CM insert gently with Milli-Q® water. Repeat twice.
7.   Add 0.5% hydrochloric acid in 70% ethanol for 45 seconds to remove excess stain.
8.   Rinse with Milli-Q® water. Repeat twice.
9.   Add diluted ammonium hydroxide and leave for approximately 45 seconds.
10.  Rinse with Milli-Q® water. Repeat twice.
11.  Store wet or mount for microscopy.

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Immunocytochemical staining is a technique employing fluorescently labeled antibody, by which cells can be localized, labeled, and examined via fluorescent microscopy.

Materials and Reagents

• Millicell® Cell Culture Inserts and Insert Plates
• Sterile Phosphate Buffer Saline (PBS)
• Methanol, 100%
• Glycerol
• FITC-conjugated antibody
• 1% BSA in PBS
• Glass slides

Method
1.   Aspirate cell culture media and wash Millicell® inserts or plates gently on both sides with PBS. Incubate for 5 minutes and repeat 2–3X. Consult the recommended working volumes table for appropriate volumes.
2.   Add fixative solution (e.g. methanol) for 5–10 minutes. It is not required to treat the underside of the membrane with fixative. Incubate according to protocol instructions.
3.   After treatment, aspirate fixative and fill filter wells with washing buffer. Repeat 2–3X in order to fully remove the fixative solution from both sides of the filter membrane. Do not allow cells to dry.
4.   Dilute primary antibody according to vendor recommendations. In order to obtain best results, it is recommended that optimal working dilutions be determined by the user. If permeabilization is required (such as for cytoplasmic or nuclear antigens), saponin can be added to the solution at a concentration of 0.1%
5.   Add antibody solution to each well then incubate at recommended temperature (typically room temperature or 4°C) with mild shaking or rocking to assure that solution wets out entire filter surface. If antibody is fluorescently labeled (direct labeling), cover plate with foil to protect from light.
6.   Aspirate antibody solution and wash both sides of membrane as indicated in Step 1 to remove all unbound antibody.
7.   If performing indirect labeling with a secondary antibody, repeat steps 4 through 6 with the secondary antibody. For visualization using fluorescent antibodies, continue to Visualization and Microscopy procedures. For enzyme linked assays (HRP, etc.), follow vendor procedures for developing.

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The Microscopic Examination of Samples Can Be Performed in Three Modes:
1.   Viewing from below the plate (through transparent PET or CM membranes)
Millicell® devices using PET or CM membrane have been designed to allow visualization of cells from below using an inverted microscope. For viewing live cells, microscopic observations can be made through the receiver or plastic plate containing media. In order to focus on the cells, the microscope objective (typically 5–20X) must have an appropriate working distance. (For objective specifications, visit the websites listed in the Microscope Objective Information section.) Fixed cells that do not require to be visualized in media can be viewed directly without the receiver plate. However, care should be taken not to contaminate the objective with liquid residue (media, mounting fluid) on the membrane.
2.   Viewing from above the plate (Millicell® 6-well inserts, Millicell®-24 cell culture insert plates)
Some cell culture platforms can allow the cells to be viewed in a conventional microscope directly from above using low magnification. Cells can be visualized through the lid to maintain sterility or with the lid removed for fixed cells or when maintenance of sterility is not required. Working distances of the objective must be longer when reading from above compared to when reading from below. If using immunofluorescence, it is recommended to use a mounting fluid that contains an anti-fade additive to prevent photobleaching.
3.   Visualizing membranes on microscope slides (for higher magnification or withobjectives with short working distances [less than 2 mm])
The membrane can be removed from each well for microscopic evaluation. This allows for higher magnification examination and storage of the slides for future use.


For visualizing from above the membrane, typically 5–20X objectives are used that have at least a 13.59 mm (A) or a 18.03 mm (B) working distance when viewing without or with the lid, respectively. For visualizing from below the membrane, 5–20X objectives are used that have at least a 2 mm (C) working distance.

To prepare membranes on microscope slide
1.   Remove the membrane from the well using a sharp scalpel to make a small incision in the edge of the membrane. Carefully cut along the inside of the well wall for approximately one quarter of the well diameter. Using forceps (Millipore cat. no. XX62 000 06), carefully hold the membrane while continuing to cut around the well diameter to remove membrane. Alternatively, a cork borer may be used to remove the membrane. Note: Use care to prevent membrane from curling.
2.   Place the membrane disk, cells facing up, onto a microscope slide.
3.   Add 50 µL mounting fluid to the membrane disk and allow it to wet out in order to prevent bubbles under the disk.
4.   Slowly lower a cover slip onto the membrane at an angle to allow air bubbles to be removed.

Microscope Objective Information
Information regarding microscope objective magnification power and working distances can be obtained from individual optical dealers or from the microscope vendors:

• Nikon Instruments: www.nikoninstruments.com
• Olympus Corporation: www.olympusamerica.com
• Carl Zeiss: www.zeiss.com

Note It is assumed that users of this procedure will be knowledgeable in TEM procedures.

Materials and Reagents

• Millicell® Cell Culture Inserts
• Phosphate Buffered Solution (PBS)
• Glutaraldehyde
• Osmium tetroxide
• Sodium cacodylate
• Sucrose, reagent grade
• Calcium chloride
• Lead nitrate
• Sodium citrate/Sodium hydroxide
• Uranyl acetate
• Ethanol
• Flat embedding trays
• Embedding Resin (i.e. EPON812)
• Fine forceps — Millipore cat. no. XX66 000 06
• Diamond Knife
• Cork Borer
• Durapore Filter Disk — Millipore

Sample Pictures


Living murine embryonic stem cell derived embryoid bodies visualized in a 1 um PET Millicell®-24 device using an Olympus IMT-2 inverted microscope


Neuron differentiation of embryonic stem cells in Millicell®-24 1 um PET filter plates. Murine embryonic stem cells were formed into suspended embryoid bodies (EBs), then transferred to 1 um PET Millicell®-24 plates for attachment and differentiation. The photo inset shows the inverted phase contrast through membrane of live EBs in the media. Neural differentiation after netinoic acid treatment of attached EBs was confirmed by anti-neurofilament immunofluoresence.

A. Processing/Cell Preparation
Note: Steps 1–5 should be done on an intact Millicell® cell culture insert or plate well.

1.   Wash cells briefly (2 times for 5 minutes each) at room temperature with phosphate buffered solution without fixative.
2.   Fix cells in 2% glutaraldehyde in 100 mm sodium cacodylate buffer, pH 7.5, at room temperature from 15 minutes to 2 hours.
3.   Wash cells (2 times for 5 minutes each) in 100 mm sodium cacodylate buffer at room temperature. Note: At this point, cells can be stored in the above buffer with 7 g sucrose/100 mL buffer at 4°C.
4.   Fix cells in 1% osmium tetroxide in either 100 mm sodium cacodylate or suitable phosphate buffer.
5.   Dehydrate cells in the following concentrations of ethanol:

Ethanol Concentration Kit (%)	Time (minutes)
30	15
50	15
70	15
95	15
100	3 x 15

Note: Dehydration of Millicell®-HA units should be performed in a metal pan that will be used as the embedding tray due to the tendency of the cellulosic membranes to be less rigid during the dehydration process. Attempts to transfer the membranes during these steps could lead to mechanical damage to the cells.

6.   For infiltration, EPON812, an EDPON substitution, or LX112 is suitable for both devices (do not use Spurr’s). The following is a general infiltration scheme:

Ethanol Concentration Kit (%)/Tray (% Plastic)	Time (minutes)
75/25	30 on a shaker
50/50	30 on a shaker
0/100	30 each/3x on a shaker
0/100	Overnight

Note: It is not necessary to use any other agent, such as propylene oxide, with plastic. Propylene oxide will dissolve the cellulosic filters. In addition, the standard inversion/rotation of specimens used in these steps is not advised since either (1) damage to the cell layer or (2) stretching of the cellulosic filter may occur. Mild shaking on a gel shaker apparatus is sufficient for successful infiltration.

Note: Before the next step the membrane must be detached from the surrounding plastic ring. Sometimes this will occur without manipulation since the EPON may loosen the membrane-to-ring bond. If this does not occur, use a sharp scalpel or a cork borer and cut the membrane. It may also help to cut the membrane over a 47 mm filter support disk. Under no circumstances should the membrane be left attached to the ring during polymerization.

7.   Transfer to fresh plastic and polymerize at 68°C overnight.

B. Sectioning Notes
1.   Nitrocellulose (HA), polycarbonate (PC) and polyethylene terepthalate (PET) membrane: These membranes can be sectioned in any plane without difficulty.
2.   CM (Biopore) membrane: The Biopore membrane must be processed in one of two ways based on the final thickness of the section.