Collagen may be used to coat glass coverslips for the growth of epithelial, endothelial and muscle cells, neurons, PC12 and CHO cell lines. Type I collagen is most often used for coating substrates for cell culture because it is easily obtainable from rat tails. For short term cultures, collagen can be simply applied to glass coverslips and allowed to dry.

1. Dilute collagen solution 1:10 – 1:50 with 30% ethanol and spread over surface of sterile glass coverslip.
2. Air dry in a tissue culture hood.
3. Cells can be seeded directly on the collagen surface.
4. Collagen coating prepared in this way tends to detach from the glass in long-term cultures.

Collagen IV is the major constituent of basement membrane and is, therefore, a more physiological coating for the culture of many cell types. For long-term cultures, collagen I and IV can be applied to glass coverslips by first coating the glass with polylysine or polyornithine. This provides a more stable collagen coating.

1. Prepare polylysine or polyornithine (MW of 30,000 – 70,000) at 0.1-1 mg/ml in 0.15 M borate buffer (pH 8.3). Filter sterilize.
2. Add enough solution to pool over surface of sterile glass coverslip.
3. Incubate 2-24 hours at room temperature.
4. Aspirate solution and wash coverslips 3 times with water.
   Pool collagen solution, 100 ug/ml in water over surface of coverslip.
5. Incubate 4 – 16 hours.
6. Rinse once with media and seed with cells.

Alternatively, for long-term cultures, double layered collagen coatings can provide a stable coating.

1. Spread a couple of drops of sterile collagen I solution on the sterile glass coverslip.
2. Immediately neutralize for 2 minutes with ammonium hydroxide vapors by placing the dish of coverslips in a covered dish containing filter paper wet with concentrated ammonium hydroxide. This will cause the collagen to gel.
3. Wash coverslips twice with sterile water.
4. Gently spread a couple of drops of collagen over the surface of the gelled collagen and air dry.
5. Use within a few hours for cell culture.

Gelatin can also be used for the culture of some cell types including glial cells.

1. Dissolve 100 mg gelatin in 100 ml water (triple glass distilled or RO).
2. Autoclave to sterilize.
3. While hot, thoroughly mix gelatin solution.
4. Add enough solution to pool over surface of sterile glass coverslip.
5. Chill for 2-24 hours at 4oC.
6. Remove gelatin by aspiration and add sterile water.
7. Dishes can be stored for up to one week at 4oC.
8. Remove water immediately before use for cell culture.

Nearly all types of cells adhere to these polymers of basic amino acids. They are particularly useful for the culture of CNS neurons. The L- or D-isomers can be used for cell attachment, however, the D-isomer may be preferred because it is not subject to breakdown by proteases released by cells.

1. Prepare polylysine or polyornithine (MW of 30,000 – 70,000) at 0.1-1 mg/ml in 0.15 M borate buffer (pH 8.3). Filter sterilize.
2. Add enough solution to pool over surface of sterile glass coverslip.
3. Incubate 2-24 hours at room temperature.
4. Aspirate solution and wash coverslips 3 times with media or PBS.
5. Immediately add cell suspension or growth media.

Fibronectin is an extracellular matrix constituent use for the culture of endothelial cells, fibroblasts, neurons and CHO cells.

1. Stock solution can be prepared by dissolving 1 mg/ml fibronectin in PBS. Filter sterilize and freeze in aliquots.
2. Diluted stock solution to 50-100 ug/ml in basal medium or PBS.
3. Add enough solution to pool over surface of sterile glass coverslip.
4. Incubate for 30-45 min at room temperature.
5. Aspirate to remove fibronectin and rinse coverslips with media or PBS.
6. Immediately add cell suspension or growth media. Do not allow coating to dry.

Laminin is an extracellular matrix constituent used for the culture of neurons, epithelial cells, leukocytes, myoblasts and CHO cells.

1. Stock solution can be prepared by dissolving 1 mg/ml laminin in PBS. Filter sterilize and freeze in aliquots.
2. Diluted stock solution to 10-100 ug/ml in basal medium or PBS.
3. Add enough solution to pool over surface of sterile glass coverslip.
4. Incubate several hours at room temperature.
5. Aspirate to remove laminin and rinse coverslips with media or PBS.
6. Immediately add cell suspension or growth media. Do not allow coating to dry.
7. Coating the glass coverslip first with polylysine or polyornithine and then laminin may increase the concentration of laminin applied using this method.

RECOMMENDED FOR LABORATORY USE ONLY

1.1    General staining of proteins deposited on porous nitrocellulose films for sample protein quantification.

2.1    Equipment

  2.1.1        Orbital shaker

  2.1.2        ArrayCAM™ Microarray Imager (Grace Bio-Labs)

2.2    Materials and Reagents

2.2.1        Glass staining dishes with lids (Wheaton Cat# 900203).

2.2.2        Glass slide racks with handles (Wheaton Cat# 900204, 900205).

2.2.3        ProPlate™ modules (Grace Bio-Labs)

2.2.4        Fast Green FCF (Sigma-Aldrich Cat#F7258)

2.2.5        Methanol

2.2.6        Glacial Acetic Acid

2.2.7        Distilled Water

2.3    Solutions

2.3.1        De-Staining Solution (1000mL):

2.3.1.1  Methanol (300mL, 30%)

2.3.1.2  Glacial Acetic Acid (70mL, 7%)

2.3.1.3  Water (630mL, 63%)

2.3.2        Fast Green Stock Solution (400x, 10mL)

2.3.2.1  Fast Green (0.1g)

2.3.2.2  De-Staining Solution (9.9mL)

2.3.3        Fast Green Staining Solution (1x, 1000mL)

2.3.3.1  Fast Green Stock Solution (2.5mL)

2.3.3.2  De-Staining Solution (997.5mL)

2.3.4        1% NaOH

3.1    Transfer slides to a staining jar containing fresh distilled water. 

3.1.1        Wash for 5 minutes with agitation (105 rpm on orbital shaker).

3.2    Transfer slides to a staining jar containing 1% NaOH.

3.2.1        Incubate for 15 minutes with agitation (105 rpm on orbital shaker).

3.3    Transfer slides to a staining jar containing fresh distilled water.

3.3.1        Rinse briefly by submerging slides repeatedly 10-20 times over 1 minute.

3.4    Transfer slides to a staining jar containing fresh distilled water.

3.4.1        Wash for 10 minutes with agitation (105 rpm on orbital shaker).

3.5    Transfer slides to a staining jar containing De-Staining Solution.

3.5.1        Wash for 15 minutes with agitation (105 rpm on orbital shaker).

3.6    Transfer slides to a staining jar containing Fast Green Staining Solution.

3.6.1        Wash for 3 minutes with agitation (105 rpm on orbital shaker).

3.7    Transfer slides to a staining jar containing fresh water.

3.7.1        Rinse briefly by submerging slides repeatedly 10-20 times over 1 minute.

3.8    Transfer slides to a staining jar containing De-Staining Solution.

3.8.1        Wash for 15 minutes with agitation (105 rpm on orbital shaker)

3.9    Transfer slides to a staining jar containing fresh water.

3.9.1        Rinse briefly by submerging slides repeatedly 10-20 times over 1 minute.

3.10  Dry the slide by centrifugation.

3.11  Scan slides using a fluorescent imager capable of measuring at 800 nm (settings and results may vary with imaging system and filters employed):

3.11.1    ArrayCAM Microarray Imager

3.11.1.1                      800 nm bandpass

3.11.1.2                      Typical Settings:

6.2.1.2.1        Exposure: 200 msec.

6.2.1.2.2        Acquisition Time: 4 sec.

6.2.1.2.3        Gain: 20%

3.12 See Appendix for typical results obtained with this method.

4.1    Loebke, et al. (2007) Infrared-based protein detection arrays for quantitative proteomics. Proteomics 7, 558-584.

4.2    Levine, et al. (2006) Quantitation of protein on gels and blots by infrared fluorescence of Coomasie blue and Fast Green. Analytical Biochemistry 350, 233- 238.

Figure 1. Fast Green Protein Quantification on ONCYTE® Film Slides with near IR detection using ArrayCam.

Figure 2. Quantitation of Cell Lysates on RPPA using Fast Green with ArrayCAM produces results comparable to Sypro Ruby Protein Stain measured on a GenePix scanner (Molecular Devices, Sunnyvale, CA).

Figure Legends:

Figure 1.  Total protein may be quantified after spotting on ONCYTE® AVID Film Slides by staining with Fast Green and measuring fluorescence using an ArrayCAM, an imaging system developed at Grace Bio-Labs.   ArrayCAM is a CCD-based imaging system using laser excitation at 405 nm. Fast Green emission is 800 nm. (NOTE: Similar results were obtained with ONCYTE® SuperNOVA Film Slides from Grace Bio-Labs.)

A.  Shown is a range of standard proteins from 8 – 500 pg after on-slide staining with Fast Green.

B.  Detection and quantitation of BSA, Lysozyme, and IgG using a Fast Green staining protocol (see attachment) with ArrayCAM are linear down to the lowest protein deposition (8 pg). 

 

Figure 2. Fluorescent signal for total protein detection with Fast Green/ArrayCAM yields linear results at cell lysate concentrations commonly used for RPPA analysis down to approximately 8 pg protein per spot.   Similar results were obtained using a standard protein stain method (SYPRO® Ruby total protein stain; 532nm/575nm detection) with the use of a focused-laser microarray scanner (Molecular Devices GenePix 4400).  Cell lysates were from Calyculin A-treated Jurkat cells spotted in 2-fold serial dilution starting with a concentration of 1 mg/ml in a Tris/SDS/Glycerol lysis buffer.