Dear Angela.  I think the gel that you have shown looks good, with an even blue colour across the whole of the gel around your samples.  There is a bit of a "wave" in the run - to reduce this you may try running the gel a bit more slowly.  The pale area at the top of the gel is not unusual - may be to do with high molecular weight materials (eg gelatinase-lipocalin complexes), and the transition into the gel from the well.  You can get really uneven background colour if you are running samples with a lot of salts in them (eg concentrated culture media), so the centre is a significantly different colour than the periphery - which make them a challenge to analyse across a gel (nothing like your gel here!).  The darker bands are abundant proteins that are staining darker with the Coomassie.  Consider when you perform your densitometric analysis - you should analyse each band as equal areas (ie. the same sized box) so they are comparable.  You can use a same sized box as a blue "no sample" control [I suggest you take 2 or 3 and take an average]), at the same level as your samples.  Good luck.

Gelatin Zymography

Gelatinase substrate gel electrophoresis was performed with the use of precast gels (10% polyacrylamide containing 0.1% gelatin, Novex).

Samples were prepared by dilution (1:200) into a loading buffer consisting of 0.4 mol/L Tris, pH 6.8, 5% SDS, 20% glycerol, 0.03% bromophenol blue, and 20 µL of serum loaded per lane.

After electrophoresis at 125 V, the gels were incubated in renaturing solution (2.5% Triton-X-100) for 30 minutes at room temperature and then for 72 hours at 37°C in a developing buffer containing 50 mmol/L Tris, pH 7.5, 200 mmol/L NaCl, 4 mmol/L CaCl2, and 0.02% Brij-35. When appropriate, the developing buffer contained 20 mmol/L EDTA, a known inhibitor of MMPs, or l mmol/L PMSF, a known inhibitor of serine proteases. The gels were then stained with Coomassie blue, and regions without staining were indicative of gelatin lysis.

Serum samples were prepared as follows for activation with APMA. First 25 µL of 5 mmol/L APMA was added to 100 µL of undiluted serum and incubated in a 37°C water bath for 90 minutes. The reaction was stopped with 10 mmol/L EDTA, and the samples were dialyzed for 18 hours to remove the APMA. The samples were then loaded on gels, and the zymography protocol was followed as above.

Zymography Protocol:

By:Dr. Ken Swanson

Zymography Protocol:

Zymography allows the detection and rapid identification of matrix metalloproteinase

(MMP) activities in biological samples. Samples containing MMP, including tumor and cell lysates, and body fluids such as cerebral spinal fluid (CSF) are subjected to separation by SDS-PAGE in a resolving gel containing a co-polymerized protease substrate such as gelatin. After being resolved in this gel, proteases are re-natured and act to hydrolyze the protein substrate in the gel matrix. The presence and relative activity of MMP’s in the sample are thus determined by a decrease in Coomassie brilliant blue staining of the digested gelatin at the position of the MMP. Since Coomassie® Brilliant Blue fluoresces in the near infrared and can readily be detected within the 700 channel of the LI-COR Odyssey, this MMP-dependent decrease in Coomassie staining can be readily detected and quantified using the protocol below. This protocol allows for a more precise determination of the relative amount of MMP activity than has previously been possible by zymography and provides a more convenient means of acquiring, storing and presenting this data.

Reagents:

Gelatin (Sigma-G8150)

Calcium Chloride(Sigma)

Tris-Acetic? Acid

Tris-Base?

Glycine

Triton X®-100

30% Acrylamide 0.8% Bis-Acrylamide?

TEMED

Ammonium Persulfate

Sodium Dodecylsulfate

Coomassie Brilliant Blue R-250 (Biorad 161-0406)

CSF zymography protocol

• 1.The resolving gel is made as follows: 40 mg of gelatin in 10 ml of deionized water (70oC). In order, add 10 ml of deionized water (21oC) to cool the mixture, 10 ml of 4X separating buffer (1.5M Tris HCl, pH 8.8) and 10 ml of 30% acrylamide 0.8% bis-acrylamide (Sigma) solution. Initiate polymerization by the addition of 200 l of TEMED (Sigma) and cast as aml of ammonium persulfate (Sigma) and 50 m mini-gel (8 cm X 5 cm X 0.75mm).

• 2. After the resolving gel is fully polymerized, pour the stacking gel (9 ml of distilled water, 3.75 ml of stacking buffer (0.5M Tris HCl, pH 6.8), 2.25 ml of 30% acrylamide, 0.8% bis-acrylamide, l TEMED) and place the comb.ml APS and the addition of 30m200 

• Denature samples containing the MMPs to be analyzed in 1X Laemmli sample buffer with BME and electrophorese within the gel described above at 25 mA until desired resolution is attained. Running buffer: 25 mM Tris pH 6.8, 190mM glycine and 3.5 mM SDS in 500 ml of distilled water.

• After electrophoresis is complete, incubate the gel in 2.5% Trition X®-100 (Sigma) washing buffer with shaking at 10 rpm for 1 hour.

• Incubate in digestion buffer (10 mM calcium chloride, 20mM Tris Acetic Acid with pH of 7.5.) for 8 to 12 hours 37°C.

• Stain the gel for an hour in 26.6ml of 3X Coomassie Brilliant Blue R-250 (Biorad) solution, (0.1% mass/vol of Coomassie Blue), with 13% acetic acid, De-stain in 10% acetic acid and 5% methanol for 3 hours.

• Scan the de-stained gels on the LI-COR Odyssey® using the 700 channel. Scan at intensity 5 as a starting point

• Quantify digested areas using the ‘Average’ background method in the Background settinngs. Since the Odyssey is detecting a net decrease in intensity, maintain the same size for each quantification shape that is drawn.

Notes:

• Optimal mass of material to be analyzed and time of digestion for each experimental paradigm must be determined empirically. Bands that are completely digested in the center diverge from linearity, thus over digestion should be avoided. It is suggested that a concentration curve be generated at the beginning of a study using the Odyssey to determine the optimal mass of the material being studied. Once established, the time and temperature of the reactions must be adhered to during subsequent experiments.

• Incompletely dissolved gelatin (step 1) and low concentration of Coomassie Brilliant Blue (step 5) may lead to uneven staining and poor quantification.

References

1. Perides G, Asher RA, Lark MW, et al. Glial hyaluronate-binding protein: a product of metalloprotease digestion of versican? Biochem J 1995; 312: 377-384.

2. Perides G, Charness ME, Tanner LM, et al. Matrix metalloproteases in the cerebrospinal fluid of patients with Lyme borreliosis. J Infect Dis 1998; 177: 401-408.

3. Friedberg MH, Glantz MJ, Klempner MS, et al. Specific matrix metalloprotease profiles in the cerebrospinal fluid correlates with the presence of malignant astrocytomas, brain metastases, and carcinomatous meningitis. Cancer 1998; 82: 923-930.

4. Timothy E, Van Meter TE, William C, et al. Induction of membrane-type-1 matrix metalloproteinase by epidermal growth factor-mediated signaling in gliomas. Neuro-Oncol? 2004; 6(3):188-199

Coomassie Staining

0.2% Coomassie Gel Stain (in 7.5% acetic acid and 40% methanol):

0.2 g Coomassie

7.5 ml acetic acid

40 ml methanol

Q.S. with H2O to 100 ml. Nalgene or Whatman filter before use.

Coomassie Rapid Destain (7.5% acetic acid and 5% methanol):

75 ml acetic Acid

50 ml methanol

Q.S. with H20 to 1 L.

Dear Angela.  I think the gel that you have shown looks good, with an even blue colour across the whole of the gel around your samples.  There is a bit of a "wave" in the run - to reduce this you may try running the gel a bit more slowly.  The pale area at the top of the gel is not unusual - may be to do with high molecular weight materials (eg gelatinase-lipocalin complexes), and the transition into the gel from the well.  You can get really uneven background colour if you are running samples with a lot of salts in them (eg concentrated culture media), so the centre is a significantly different colour than the periphery - which make them a challenge to analyse across a gel (nothing like your gel here!).  The darker bands are abundant proteins that are staining darker with the Coomassie.  Consider when you perform your densitometric analysis - you should analyse each band as equal areas (ie. the same sized box) so they are comparable.  You can use a same sized box as a blue "no sample" control [I suggest you take 2 or 3 and take an average]), at the same level as your samples.  Good luck.

I agree with Simon; that's not a bad zymograph.  If you'd run it out a little longer, it would likely have straightened out more 

Observations

1. To improve the running of the gel , I recommend a) make a run without a sample for 10 minutes ; b ) preparing the running buffer and the gels again ; c ) have only 5 lane samples, however in the sixth lane (right to left ) is contamination of the 5 lane .

2. lysis bands are proteins that are degrading the substrate , the dark bands are other proteases that degrade other substrate and that's interesting.

I hope you find them useful and thank you very much for contacting me

Hector Flores- Herrera

Dear Eloy,

      I advise you to utilize new electrophoresis buffer before conducting the electrophoresis. According to my own experience, if I use the electrophoresis buffer for more than three times, the quality of zymography is likely to be poor.

     Hope my suggestion will be helpful.

     Sincerely

                                                                                           Kaizhou Yan

Dear Eloy,

I agree with Simon and Kaizhou. Your gel is not so bad. In your case, I will dilute the sample to be sure that all gelatinolytic bands are not saturated. For each electrophoresis, I change the different buffers (electrophoresis, triton and incubation) to be sure that I do not have any contaminations like bacteria. In your gel, you have a continous white band in all samples (molecular weight marker included). It's probably this sort of contamination.

You can also stain your gel with G-250 coomassie blue (more sensitive) and with a better background.

Did you solubilized your gelatin solution 30minutes at 37°C before use?

Sincerely,

Dear Eloy,

Your zymogram is not bad at all.

Dear Angela,

I agree. It looks quite good!