Protein Stains Support — Getting Started

You can use the SimplyBlue™ SafeStain for protein quantitation by densitometry. However, the Colloidal Blue Staining Kit (Cat. No. LC6025) is the best suited for this application.

For gels that you are pouring yourself, add Rhinohide™ Polyacrylamide Gel Strengthener (Cat. Nos. R33400, R33410) to the formulation. Decrease the shaker speed/motion; only very gentle rocking is required. Slowly decant solutions during solution changes and avoid touching the gel as much as possible when pouring out solutions.

We recommend storing the SimplyBlue™ SafeStain at room temperature where it is stable for six months.

SimplyBlue™ SafeStain is a ready-to-use proprietary staining solution that contains Coomassie™ G50. It does not contain acetic acid or methanol.

With the regular staining protocol, 7 ng of BSA can be detected after destaining whereas with the microwave staining protocol, 5 ng of BSA can be detected.

SimplyBlue™ SafeStain can be used to stain dry PVDF membranes. Staining nitrocellulose and wet PVDF membranes results in high background and is not recommended.

Proteins stained using the SimplyBlue™ SafeStain are compatible with mass spectrometry (MS) analysis.

SimplyBlue™ SafeStain does not contain acetic acid or methanol. An alcohol/acetic acid fixing step prior to staining with SimplyBlue™ SafeStain is not required or recommended.

SimplyBlue™ Safestain is specially designed for safe, non-hazardous disposal. It does not contain methanol or acetic acid and does not require the use of methanol or acetic acid which must be disposed of as hazardous waste.

Coomassie™ G250 is more sensitive than Coomassie™ R250 whereas Coomassie™ R250 stains faster than Coomassie™ G250.

Our recommendation would be to separate the protein from the Coomassie™ G250 stain by electroelution or alcohol treatment so as to not illicit an immune response to the Coomassie™ G250. If the antibodies are to be used in membrane-based immunodetection, then inserting the membrane-bound band subcutaneously can produce antibodies that work especially well in membrane-based detection.

There should be no problem with using a SimplyBlue™ stained protein band for immunization. The process of staining and destaining the gel should remove SDS (if an SDS PAGE gel is used) so it will not interact with the Freund's Adjuvant.

We recommend leaving the gel in the stain overnight. There is no need to add the extra salt solution as recommended in the manual because there are no buffer salts and SDS to worry about, and that would end up inhibiting the staining too much.

You can use SimplyBlue™ SafeStain to stain the PVDF membrane to detect your protein to be sequenced. After detection and prior to sequencing, rinse the membrane in 20–30% ethanol to remove the remaining stain.

We recommend storing the Imperial Protein Stain at room temperature where it is stable for a year.

Imperial™ Protein Stain contains Coomassie R-250. The rest of the composition is proprietary.

With the enhanced protocol, the Imperial™ Protein Stain can detect less than 3 ng protein per band in 3 hours.

Yes, here is the procedure for processing gels stained with Imperial ™Stain, for mass spectrometry analysis.

We recommend storing the PageBlue™ Protein Staining Solution at room temperature where it is stable for a year.

PageBlue™ Protein Staining Solution contains Coomassie G-250 for protein staining on polyacrylamide gels and PVDF membranes. The rest of the composition is proprietary.

The PageBlue™ Protein Staining Solution can be reused up to three times without noticeable loss in detection sensitivity.

It can detect proteins with a dynamic range of 5–500 ng, which is ~10 times more sensitive than traditional Coomassie R-250-based dyes.

Fixing gel proteins with 25% isopropanol/10% acetic acid solution or 12% trichloroacetic acid (TCA) for 15 minutes can increase staining sensitivity.

The first wash step in the staining procedure is designed to remove sodium dodecyl sulfate (SDS) from the gel, because SDS interferes with staining. Native gels do not contain SDS and, therefore, this step can be omitted from native PAGE applications.

Small proteins (< 10 kDa) are susceptible to leaching from the gel during the staining procedure and require fixation with glutaraldehyde before staining the gel with PageBlue™ Protein Staining Solution. Other common protein fixatives (e.g., acetic acid, isopropanol, ethanol, TCA, etc.) are not suitable for this purpose, as proteins will be washed out of the gel during the staining procedure. Please see procedure for fixation, mentioned on Page 3 of the manual.

We recommend storing the Colloidal Blue Staining kit at room temperature where it is stable for 1 year.

The Colloidal Blue Stain is five times more sensitive than traditional Coomassie™ Blue staining techniques. Using the Colloidal Blue Staining Kit, <10 ng of BSA can be detected on a 4–20% 1.0 mm Novex™ Tris-Glycine Gel in 1 hour. Non-reduced samples stain slightly more intensely than reduced samples. Bands are visible after 1 hour in staining solution.

Proteins stained using Colloidal Blue Stain are compatible with mass spectrometry (MS) analysis.

No we do not recommend staining membranes with the Colloidal Blue (G-250) Staining Kit as the background will be too high. Better alternatives include:

1. Coomassie™ (non-colloidal) staining: Stain in 0.1% Coomassie™ Blue R-250 in 50% methanol for 5 minutes and destain with several changes of 50% methanol and 10% acetic acid. Rinse with several changes of water, air dry and store for up to 12 months at –20 degrees C. Sensitivity is approximately at the 50–100 ng level.
2. SimplyBlue™ SafeStain. There is a protocol included in the SimplyBlue™ SafeStain manual for staining PVDF but it is not recommended for nitrocellulose because of the high background.
3. Amido Black: same as Coomassie™ but less sensitive.
4. Ponceau S: same as Coomassie™ but less sensitive.
5. UV transillumination: After blotting, place the membrane on a filter paper and allow to dry at room temperature for about 10 minutes. Rewet in 20% methanol and view the blot in front of white light while it is still wet; the bands will look more translucent than the membrane. If the bands disappear because the membrane is dry, rewet the membrane.
6. Novex Reversible Membrane Protein Stain (Cat# IB7710): Allows for complete, reversible staining of protein on nitrocellulose & PVDF membranes. Sensitivity is higher than Ponceau S (<10 ng of BSA in 10 mins as blue bands) and the staining is reversible in 5 minutes. Western blot detection is unaffected by the staining and erasing process, and in some cases higher sensitivity is achieved.

Yes, it stains non-reduced proteins more intensely.

The Colloidal Blue Staining Kit uses Coomassie™ G-250 whereas standard Coomassie™ staining protocols use Coomassie™ R-250. Coomassie™ G-250 is a turquoise color whereas Coomassie™ R-250 is closer to navy blue. The difference is in the shade of color, rather than the intensity.

No, due to its chemistry, the Colloidal Blue Staining Kit stains only proteins and peptides. The SilverXpress™ Staining Kit is a highly sensitive stain for nucleic acids.

• Do not leave the stained gel in pretreatment gel drying solutions, such as Gel-Dry™ Solution, for more than 5 minutes.
• Prolonged exposure to pre-treatment gel drying solutions will destain the gel completely.

A 5% bleach solution will effectively remove the Colloidal Blue stain from plastic, porcelain and metal surfaces.

The Colloidal Blue Staining kit can be stored in the refrigerator, but be sure to let the kit return to room temperature before use. Also remember to shake the Stainer B reagent prior to use. Cold storage will not enhance the shelf life of the stain.

Yes, the fix serves two purposes: it fixes the sample in the gel and it helps to remove gel background. If you do not use the fixing solution, the background on the gels will be high and detection will be less sensitive. High background can be caused by ampholytes remaining in the gel.

No, the staining solution should be prepared fresh, no more than 24 hours before use. If fresh solution is not used, staining background will be high.

No, heat causes colloids to solubilize; this in turn, causes the free-dye in solution to increase which causes an increase in background and a decrease in staining intensity.

Colloidal Blue Coomassie™ G250 Stain requires a wavelength of 610 nm whereas standard Coomassie™ R250 stains require a wavelength of 588 nm.

Some of the components of the Colloidal Blue Stain are probably too harsh to be used in conjunction with protein sequencing, although the actual Coomassie™ dye itself does not have an adverse effect.

Yes, Colloidal Blue Stain can be used before western blotting. However, for optimal transfer efficiency, we recommend destaining the gel and then equilibrating in a series of Tris base/Glycine/SDS solutions to increase solubility; when the transfer is complete, the membrane should be treated with methanol to remove the stain prior to chromogenic development (not necessary prior to chemiluminescent detection).

Coomassie™ G250 is more sensitive than Coomassie™ R250 whereas Coomassie™ R250 stains faster than Coomassie™ G250.

Yes, the Colloidal Blue Staining kit works very well with Novex™ Zymogram gels.

We recommend storing the Pierce™ Silver Stain for Mass Spectrometry at room temperature where it is stable for a year.

The Pierce™ Silver Stain for Mass Spectrometry provides subnanogram sensitivity, detecting proteins at less than 0.25 ng per band in 30 mins after fixing.

After using the Pierce™ Silver Stain for Mass Spectrometry, proteins can be recovered and processed by several methods for mass spec analysis. For in-gel trypsin digestion, with or without reduction and alkylation, use the Thermo Scientific™ In-Gel Tryptic Digest Kit (Cat No. 89871). Consider following the recommendations of the core facility that will be performing the mass spec analysis.

We recommend storing the Pierce™ Silver Stain Kit at room temperature where it is stable for a year.

The Pierce™ Silver Stain Kit detects proteins at 0.25 ng per band.

The gel is washed and fixed to remove electrophoresis buffer salts. The silver stain is added, releasing silver ions that interact with the proteins. The developer is then added causing the protein bands to darken as the bound silver reduces. The color development is stopped by lowering the pH with acetic acid.

The stain performs very well for most polyacrylamide gel types (i.e., suppliers and buffer systems). Both 1D and 2D polyacrylamide gels can be stained with this kit. The default protocol is optimized for gels that are 1.0mm thick; incubation and wash times may require adjustment to achieve optimal results for gels of other thicknesses.

A detection level of 0.4 ng per protein band has been achieved. Most proteins are easily detectable at low-nanogram (1–5 ng) amounts.

Yes, see here for the procedure for processing gels stained with Pierce™ Silver Stain, for mass spectrometry analysis. However, for best results, we recommend using the Pierce™ Silver Stain for Mass Spectrometry (Cat. No. 24600). This kit includes an optimized procedure and all the necessary reagents for staining gels and destaining gel pieces before in-gel trypsin digestion and mass spectrometry.

Yes. First destain the Coomassie-stained gel to completely remove background. If an acid or methanol destaining solution was used, thoroughly wash the gel with ultrapure water, then proceed with the Pierce™ Silver Stain Kit protocol.

The protocol is quite flexible with regard to fixing and staining times. After gels are fixed for at least 30 minutes, they can be stored in water until the next day. The gel-staining time can range from 5 minutes to 24 hours without incurring additional background.

Yes. Thoroughly wash the gel with water to remove the acetic acid and then soak the gel in a solution containing 5% glycerol and 10% methanol (or 10% ethanol) for one hour before drying in a gel-drying apparatus.

We recommend storing the SilverQuest™ Silver Staining kit at room temperature and the SilverXpress™ Silver Staining kit at 4 degrees C. Both kits are stable for six months when properly stored.

We recommend using the Mark12™ Unstained Standard with the SilverXpress™ and SilverQuest™ Silver Staining kits. For the SilverQuest™ Silver Staining kit, we recommend diluting the Mark12™ Unstained Standard to 1:10 and loading 5 μL per lane. For the SilverXpress™ Silver Staining kit, we recommend diluting the Mark12™ Unstained Standard to 1:20 and loading 5 μL per lane.

Yes this is possible. We recommend destaining the gel with water (it is not necessary to remove all the stain, but if you would like to do so, we recommend soaking the gel in 50% ethanol followed by numerous water washes). If the SimplyBlue™ stained gel was destained using salt, we recommend washing the gel numerous times in water to remove all the salts before proceeding with the Silver staining protocol. Further, we recommend beginning the silver staining protocol with the fix step (this will help to remove any methanol/ethanol and salts from the previous staining).

Glutamic acid, aspartic acid, and cysteine thiols are the most reactive with the SilverXpress™ and SilverQuest™ staining methods.

Yes, the SilverXpress™ staining solutions can be prepared before staining but not more than 24 hours in advance.

Note: The Fixing Solution can be stored for a month. If the solution turns pink, it needs to be remade.

The gel can be stored in the fixative overnight if there is not enough time to complete the staining protocol. Longer fixing times may improve the sensitivity and background staining in some cases.

Yes, gels can be left overnight in the second Sensitizing Solution. Although some other kits recommend leaving gels in the fix step, we have found that overnight fixation diminishes staining performance.

No, both ways have been tested in-house: adding all three ingredients at once and adding the water after combining the two stainers. Both methods give equivalent results.

We have used a slight modification that has yielded good results. We recommend skipping the fixation step and changing the composition of the sensitizer solution to 2 mL of sensitizer and 198 mL of ultrapure water. This modification gives 0.3 ng sensitivity down to 50 bases, whereas the normal protocol gives 0.9 ng sensitivity down to 50 bases and ethidium bromide's sensitivity is approximately 10 ng down to 50 bases. Further, to increase sensitivity, we recommend using 7.5 mL of Stainer A instead of 5 mL or eliminate the second wash step (which will also increase background).

The exact level of sensitivity achievable with the SilverXpress™ Silver Staining kit depends largely on the type of protein and its preparation. Typically, non-reduced samples yield excellent results in the sub-nanogram range. A 0.86 ng load of non-reduced BSA can be detected with the SilverXpress™ kit, whereas a standard Coomassie™ staining procedure can only detect above 50 ng of the same type of protein. If you are using the Mark12™ Standard, a 1:20 dilution is an appropriate dilution to use as a control. Reduced proteins may yield less sensitive results. If BME is used as a reducing agent, sensitivities equal to those of non-reduced samples can be expected. However, if, DTT is used as a reducing agent, one can expect sensitivities in the nanogram rather than sub-nanogram range.

The kit should be able to detect ≥0.3 ng of protein.

We recommend leaving the gel overnight in the second sensitizing wash solution. Although some other kits recommend leaving gels in the fix step, we have found that overnight fixation diminishes staining performance.

The SilverQuest™ Silver Staining Kit is compatible with mass spectrometry (MS) analysis whereas the SilverXpress Silver Staining kit is not compatible with mass spectrometry analysis.

Yes, due to the lack of glutaraldehyde in the sensitizing solution, you should be able to transfer the proteins to a membrane after completely destaining the SilverQuest™ stained gel.

SYPRO™ Ruby Protein Gel Stain binds primarily to proteins through ionic charges of the dye, with basic side chains (lysine, arginine, histidine and to a lesser extent with tyrosine and tryptophan. The fixative solution and SYPRO™ Ruby stain solution both have an acidic pH, and SYPRO™ Ruby dye binding increases with protonated basic residues. SYPRO™ Ruby dye will also bind SDS bound to the proteins and in the gel matrix. The high 50% methanol concentration in the fixative solution is better at stripping out the SDS from the gel matrix, lowering the background staining and allowing for an optimal signal to noise.

Here is a reference on amino acid specificity of SYPRO™ Ruby Protein Gel Stain:

Ultrasensitive fluorescence protein detection in isoelectric focusing gels using a ruthenium metal chelate stain. Steinberg TH, Chernokalskaya E, Berggren K, Lopez MF, Diwu Z, Haugland RP, Patton WF. Electrophoresis 2006, 21, 486–496.

Since SYPRO™ Ruby Protein Gel Stain binds primarily to basic protein residues instead of SDS, it can be used as an endpoint stain (direct, saturation binding with a low off-rate) whose performance is not affected by deviations in fixation, staining or destaining solution volumes or incubation times, but just the amount of protein that is present. The strong 50% methanol/7% acetic acid fixation in the SYPRO™ Ruby stain protocol removes most SDS bound to proteins and in the gel matrix, so that the signal is due to direct protein binding. Stains that bind protein indirectly via SDS/protein association (such as Deep Purple, Lucy, SYPRO™ Orange/Red/Tangerine, Nile Red dyes) show higher protein to protein and gel to gel variation depending on how much SDS is bound to the protein and present in the gel matrix. Proteins that do not bind SDS to saturation, such as heavily glycosylated or phosphorylated proteins will show reduced staining with indirect SDS binding protein stains.

Figure legend: A mixture of glycosylated (G), phosphorylated (P), and nonglycosylated/phosphorylated proteins, all at 125 ng/band were separated on 4–12% Bis-Tris gels and stained according to the manufacturer’s recommended protocols. Gels were imaged at the best excitation/emission settings for each stain. Deep Purple poorly stained the glycoprotein, glucose oxidase, and the acidic phosphoprotein, pepsin. Lucy poorly stained a1 acid glycoprotein.

Here are some references:

• Comparative performance of fluorescent total protein stains on one- and two-dimensional PAGE gels. Hart C, Ahnert N, Hajivandi M, Lindsey A, Harwood SH. Poster, Journal of Biomolecular Techniques, Volume 17, Issue 1, March 2006.
• Relative photostability and differential staining of proteins in two dimensional gels. Smejkal GB, Robinson MH, Lazarev A. Electrophoresis 2004, 25, 2511–2519.

Protein stains for proteomic applications: Which, when, why? Miller I, Crawford J, Gianazza E, Proteomics 2006, 6, 5385–5408.

Fluorescence intensity varies linearly with protein quantity over nearly three orders of magnitude, from 1 ng to 1000 ng. SYPRO™ Ruby stain has a broader linear range than silver staining, basically, the more protein you have, the more SYPRO™ Ruby stain that binds.

If a protein has a low content of basic residues, you will see a lower level of staining relative to a protein with a higher content of basic residues. Very heavily glycosylated proteins are an example of proteins with low content of basic residues and may not stain as well as non-glycosylated or lightly glycosylated proteins, thus lowering detection sensitivity. If a protein does not have any basic residues, as can exist for synthetic constructs, it will not bind the dye.

Detection sensitivity is only slightly diminished by dilution of SYPRO™ Ruby Gel Stain in water up to 5-fold. However, both the fluorescence signal as well as the dynamic range are both reduced significantly with even a 1/2 dilution. Detection sensitivity also remains high if the stain is reused up to two times, but signal intensity is reduced up to 2.5-fold in twice-used stain. Increasing the staining volume to 100 mL is recommended when reusing the stain.

Reference: Ahnert N, Patton WF, Schulenberg B. Optimized conditions for diluting and reusing a fluorescent protein gel stain. Electrophoresis 2004, 25, 2506–2510.

Proteins stained with SYPRO™ Ruby Protein Gel Stain, or any gel stain where the proteins are fixed in the gel, cannot be blotted onto membranes. Both the fixation step and the low pH of the SYPRO™ Ruby Gel Stain solution precipitate proteins into the gel matrix to prevent diffusion during staining, and thus also efficient transfer onto membranes. We recommend using SYPRO™ Tangerine Protein Gel Stain, which is a neutral pH, simple saline solution-based, non-fixative gel stain to prestain proteins before transfer.

No. Loading solutions contain so much SDS that SYPRO™ Ruby, SYPRO™ Orange and SYPRO™ Red dyes simply localize in the free SDS and bind very little of the proteins. Proteins can be covalently pre-labeled with ATTO-TAG™ CBQCA (Cat. No. A2333), DDAO succinimidyl ester (Cat. No. C34553) or TAMRA-succinimidyl ester (Cat. No. C2211) dyes, or the TC-FLAsH™ Expression Analysis Detection Kits (Cat. No. A10067 for orange fluorescence, Cat. No. A10068 for red fluorescence) prior to electrophoresis without affecting protein migration through the gel.

SYPRO™ Orange or SYPRO™ Red Protein Gel Stain can be diluted 5000-fold into the cathode (upper) buffer tank to stain proteins during electrophoresis without affecting migration. The problem with doing this is that there is considerable background fluorescence in the gels from the dye interacting with SDS. This can be reduced after electrophoresis by destaining in 7.5% acetic acid for 15–60 minutes. This method also results in poorer protein sensitivity than the standard post-staining method, requires the same amount of time before the gel can be imaged, and contaminates the electrophoresis apparatus.

Yes, but ethanol/acetic acid will produce ethyl acetate, which has a strong odor, so you should do this fixation in a fume hood.

No, other commonly used gel fixatives can be used, including reducing the methanol concentration to 10% methanol in 7% acetic acid. SYPRO™ Ruby stain itself will fix proteins in the gel and there is no need for a separate fixation step to stain proteins with reasonably good results. The 50% methanol/7% acetic acid fixation recommended in the protocol has been determined to best remove residual SDS from the gel matrix and thus give the lowest background and optimal sensitivity compared to other fixation methods. Reduced methanol concentrations could result in a heavily stained SDS front at the bottom of the gel, which will reduce detection sensitivity for low molecular weight proteins running near this region.

No, 23 min happens to be the time that is left over from the total 30 min stain time after subtracting the microwave and 5 min incubation times. The reality is that those are the minimum times used so that the whole fix, stain, destain procedure can be complete in 90 min. The exact timing of the microwave staining step 2.2 does not need to be followed stringently. Gels can be stained longer than 30 min, but background will also gradually increase along with the protein signal, so that sensitivity is not improved by staining longer than 30 min using the microwave protocol. Generally, staining for 30 to 90 min will give similar results. The maximum end-point signal intensity is reached after about 5 h, the same as that achieved using the overnight staining protocol.

Gels just need to be microwaved twice during the 30 min stain time to near boiling, so that only small bubbles are formed. Over-boiling can bump the staining solution, and in some cases, scorch and damage the gel.

Yes, the best place to stop for the day is during the first fixation step. Gels can be left overnight up to several days in the first fixation solution with no effect on the resulting staining. One long fixation is sufficient to remove SDS from the gel, so it is not necessary to repeat the fixation step, thus reducing solvent usage. An overnight or longer fixation will dehydrate the gel matrix significantly, so that it will be reduced in size and turn opaque white. You must rehydrate the gel before microwaving. Simply rock for about 5 min in water or SYPRO™ Ruby Gel Stain to rehydrate the gel. Check to make sure that the gel is floating in solution and has not stuck to the bottom of the staining dish before microwaving, or it will rehydrate unevenly and stain unevenly. After the gel has rehydrated back to its original size, it can then be microwaved in the SYPRO™ Ruby Gel Stain.

A handy list of instruments that work well with SYPRO™ Ruby stain is available here.

For many routine gel or blot images that do not require analysis, a simple digital camera or camera phone, such as an iphone camera, can be used in combination with a UV or blue light transilluminator, such as the Safe Imager™ 2.0 Blue-Light Transilluminator (Cat. No. G6600). Place the gel on the surface of the transilluminator and cover with the amber plastic filter. Take a picture through the amber plastic to reduce background illumination and improve sensitivity. Blots are best imaged from above the membrane (epiillumination) rather than through the membrane (transillumination). Place the light box on its side with the blot face-up on the table. Hold the amber plastic filter up close to the camera lens to take a picture.

SYPRO™ Ruby dye (as well as SYPRO™ Orange, Coomassie™ Fluor Orange and the nucleic acid stains SYBR™ Safe, SYBR™ Gold, and SYBR™ Green I and II) fluoresces nicely on the Safe Imager™ 2.0 Blue-Light Transilluminator (Cat. No. G6600) or other similar blue light transilluminators with excitation near 470 nm. The gel can be viewed with amber glasses that are supplied with the unit. UV light transilluminators equipped with 302 nm or 365 nm bulbs can be used as well, but would require UV-protective equipment during use.

The SYPRO™ Ruby stain does not interfere with mass spectrometry. After staining with SYPRO™ Ruby stain, the protein band or spot can be trypsinized and sent for mass spectrometry analysis. Although the SYPRO™ Ruby stain does not interfere with mass spectrometry, occasionally we see some peaks that are due to the SYPRO™ Ruby dye. These are a symmetrical grouping of peaks centering around 1257 and 1279 MW.

Reference: Parker K, Garrels J, Hines W, Butler E, McKee A, Patterson D, Martin S. Electrophoresis 1998, 19, 1920–1932.

We have only tested proteins as small as 6,000 Da, but if smaller proteins can be resolved in the gel and contain lysine, arginine or histidine basic amino acid residues, then it is likely that they will be detected too.

SYPRO™ Ruby stained gels can be post-stained with any silver stain, such as SilverXpress™ Silver Staining Kit (Cat. No. LC6100) and SilverQuest™ Silver Staining Kit (Cat. No. LC6070), and double staining is actually an excellent method to enhance the sensitivity obtained with individually stained SYPRO™ Ruby or silver stained gels. Simply follow the normal 10% methanol, 7% acetic acid destain after SYPRO™ Ruby staining, wash in water, if desired, to image the SYPRO™ Ruby signal and then perform the silver stain procedure. It is not necessary to perform the fixation step in the silver stain protocol, as the gels have already been fixed. The silver ions are apparently attracted to the SYPRO™ Ruby dye, enhancing the silver deposition around the proteins and thus the signal. Once a SYPRO™ Ruby stained gel has been silver stained, the SYPRO™ Ruby fluorescence signal can no longer be detected.

Yes, SYPRO™ Ruby stained gels and blots can be stained with any Coomassie™ Blue dye–based stain and will yield similar results as a gel or blot stained only with Coomassie™ Blue dye. The fluorescence of SYPRO™ Ruby will be lost after Coomassie™ Blue staining.

No, the SYPRO™ Ruby Protein Gel Stain and Blot Stain have very different formulations that are optimized for their respective usage and are not interchangeable. Blots or gels stained with the alternate SYPRO™ Ruby product will have suboptimal staining intensities and high backgrounds.

SYPRO™ Ruby Gel Stain is an endpoint stain, which means that once it has reached saturation binding of proteins, it will no longer continue to stain proteins or increase background signal, unlike silver stains. Gels can be left in SYPRO™ Ruby Gel Stain for months (4 degrees C recommended) without over-staining.

SYPRO™ Ruby Gel Stain has a very slow off-rate. In fact, it is very difficult to completely strip the dye out of the gel once it has bound proteins. Stained gels can be stored at 4 degrees C, protected from light, for at least several months with little loss in signal. Just keep hydrated in a little water with 2–5 mM sodium azide as a preservative or in 7% acetic acid. For more permanent storage, dry the gel or vacuum seal in 1–5 mL of SYPRO™ Ruby stain containing 2–5 mM sodium azide and store at 4 degrees C. Seal-A-Meal™ food storage bags are useful for this method of preservation.

The gel would need to be sequentially stained and imaged after each stain. The order of staining would be:

InVision™ His-Tag In-Gel Stain (Cat. No. LC6030) → Image→ Pro-Q™ Diamond Phosphoprotein Gel Stain → Image → Pro-Q™ Emerald 300 or 488 Glycoprotein Gel Stain → SYPRO™ Ruby Protein Gel Stain → Coomassie™ Blue or Silver Stain → Image

With the Safe Imager™ 2.0 Blue-Light Transilluminator, it takes 8–10 minutes of constant ultraviolet illumination to cause excessive photobleaching. The bleaching rate will vary with the intensity of your light source, but under most conditions this probably will not be a problem. Even when bleached, gels can be prestained with SYPRO™ stains with only a small decrease in sensitivity.

Yes, you may use SYPRO™ Orange Protein Gel Stain with Novex™ gels but the protocol would have to be modified. The stain is mainly to be used with 0.05% SDS running buffer while all our buffers contain 0.1% SDS. This results in high background as the SDS binds tightly to the stain. The following protocol is recommended for best results with Novex™ gels:

Wash solution: 7.5% (v/v) Acetic acid

Staining solution: 1:5000 SYPRO™ Orange in Wash solution

1. After the gel run is over, wash the gel in 100 ml of Wash solution for 10 minutes.
2. Place gel in Staining solution for up to 24 h in the dark.
3. Remove the gel from the Staining solution and rinse briefly in 100 ml Wash solution.
4. The gel is then ready to be photographed.

Gels stained with SYPRO™ dyes can be dried between sheets of cellophane, although there is sometimes a slight decrease in sensitivity. If the gels are dried onto paper, the light will scatter and the sensitivity will decrease. Other plastics are not recommended, as the plastic typically used is not transparent to UV light.

No. Dye concentrations higher than 1X do not give better detection. Instead, background fluorescence increases and, as the dye concentration increases, the dye becomes self-quenching and the signal actually decreases.

No, not completely. You can incubate the gel or PVDF blot in several changes of 20–50% methanol, 150 mM Tris, pH 9 to remove the majority of the dye, but it will not be completely gone.

Yes, the 15 min incubation time is a minimum time. The blot will not overstain if left in SYPRO™ Ruby Blot Stain longer than 15 min.

Yes. Blots can be left in water indefinitely without destaining the stained proteins. Longer water washes can help to reduce high non-specific background, especially for immersion-stained PVDF membranes.

Yes. We recommend staining with SYPRO™ Ruby Blot Stain after staining with Pro-Q™ Emerald 300 or 488 Glycoprotein Blot Stain.

Yes. We recommend staining with SYPRO™ Ruby Blot Stain after staining with Pro-Q™ Diamond Phosphoprotein Blot Stain.

Yes, staining a dry PVDF membrane (face staining) gives a lower non-specific background signal and results in a better signal to noise compared to a blot that has been re-wet in methanol (immersion staining). Background is also lower because the back side of the membrane does not see stain. Areas of the blot that are not completely dry or have a lot of residual SDS will wet out and show up as a darker stained background. If this is a problem, the entire PVDF blot can be re-wet in 100% methanol, so that the entire background is stained the same. Both the stained protein signal and blot background will be brighter. You should also increase the number of water washes or time for immersion stained blots. Drying the blot also completely binds the proteins to the membrane.

Dried blots stained with SYPRO™ Ruby blot stain are very photostable and can be stored for long periods of time at room temperature, in the dark, including taping into a notebook.

Yes. SYPRO™ Ruby stain is compatible with subsequent western detection using colorimetric, fluorogenic, and chemiluminescent detection techniques including BCIP/NBT, ECL, and CDP-Star™ reagent. For best western detection sensitivity, we recommend destaining the blot in 20% methanol, 150 mM Tris, pH 8.8 for 10 min followed by four 1 min rinses in deionized water to remove most of the SYPRO™ Ruby dye before performing the western detection procedure.

SYPRO™ Ruby stain is actually brighter when dry, so a dry blot is better for epiillumination. If you only have a transilluminator, blots are more transparent when wet, so wet illumination will give a brighter signal and lower background.

No. SYPRO™ Ruby Protein Blot Stain is not compatible with cationic membranes, such as Immobilon™ CD or Immobilon™ N membranes. Since it is an anionic dye, it binds non-specifically to the membrane. This is also true for amido black, Coomassie™ Blue, ponceau red and just about any other total protein blot stain. The Immobilon™ P or Immobilon™ FL membranes are recommended for use with SYPRO™ Ruby Protein Blot Stain.

We recommend storing the MemCode™ Reversible Protein Stain Kit for Nitrocellulose Membranes at room temperature where it is stable for a year.

The MemCode™ Reversible Protein Stain Kit for Nitrocellulose Membranes detects as low as 25 to 50 ng protein per band.

We recommend storing the Novex™ Reversible Membrane Protein Stain Kit at room temperature where it is stable for six months.

The stain is able to detect <10 ng bovine serum albumin.

The composition of the solutions in the Novex™ Reversible Membrane Protein Stain Kit is proprietary.

Novex™ Reversible Membrane Protein Stain is more sensitive than ponceau S. Ponceau S does not give a good measure of low-abundance protein transfer, or of the resolution of the separation.

The dye that is bound to the proteins in the pre-stained ladder is charged and covalently bound, so the transfer efficiency of pre-stained ladders is almost always better than that of SDS-denatured proteins. Therefore, pre-stained ladders are not a good measure for transfer success.

We do not recommend re-using the Novex™ Reversible Membrane Protein Stain solutions as this can lead to formation of a precipitate and will affect the staining.

The stain is completely reversible by using the Eraser solution and hence does not affect the immunodetection process.

We recommend storing the InVision™ His-Tag In-Gel Stain at room temperature where it is stable for 6 months.

InVision™ His-tag In-gel Stain is a ready-to-use, proprietary fluorescent stain that is specially formulated for fast, sensitive, and specific detection of His-tagged fusion proteins. It consists of a proprietary fluorescent dye conjugated to Ni2+: nitrilotriacetic acid (NTA) complex. The Ni2+ binds specifically to the oligohisitidine domain of the His-tagged fusion protein allowing specific detection of His-tagged fusion proteins from a mixture of endogenous proteins.

The InVision™ His-Tag In-Gel Stain is pink in color.

The stain is capable of detecting ~0.5 pmole of a 6X His-tagged fusion protein (e.g., 1 pmole of a 30 kDa protein is 30 ng). The staining intensity is sensitive to the number of moles of protein contained in a protein band as 1 molecule of InVision™ His-tag In-Gel Stain binds to only 1 oligohistidine tag molecule of the protein. For example, if you load 150 ng/band of 2 proteins with a molecular weight of 150 kDa and 30 kDa, respectively, after staining with InVision™ His-tag In-Gel Stain, the 30 kDa band stains more intensely than the 150 kDa band. This is because there is only 1 pmole of the 150 kDa band while there are 5 pmoles of the 30 kDa band in the total mass loaded (150 ng/band). The staining of a mini gel is complete in less than 3 hours.

The maximum excitation wavelength for InVision™ His-tag In-gel Stain is at 560 nm and maximum emission wavelength is at 590 nm.

To visualize His-tagged fusion protein bands after staining, you will need a:

• UV transilluminator (302 nm) equipped with a camera capable of integration - to view and photograph a gel on the UV transilluminator, use a standard video camera, CCD (charged couple device) camera, or a cooled CCD camera with ethidium bromide filter or band pass filter encompassing the emission maxima (590 nm) of the stain. A Polaroid™ camera is not recommended. Note: You can use 365 nm UV transilluminator, but you may have to expose the gel for a longer time, as the sensitivity is lower than using 302 nm UV transillumination.
  or
• Laser-based scanner with a laser line that falls within the excitation maxima of the stain (560 nm), and a 560 nm long pass filter or a band pass filter centered near the emission maxima of 590 nm. The sensitivity of detection is 2-fold more with laser-based scanners than with UV transillumination.

The BenchMark™ His-tagged Protein Standard is ideal for use as a positive control for the InVision™ His-tag In-gel Stain. The standard is formulated to allow simultaneous detection of standard proteins and His-tagged fusion proteins. The BenchMark™ His-tagged Protein Standard is included in the InVision™ His-tag In-Gel Staining Kit (Cat. No. LC6033).

The manual has a protocol for staining His-tagged fusion proteins transferred onto a nitrocellulose membrane. This procedure is not recommended for staining PVDF membranes.

Proteins stained with InVision™ His-Tag In-Gel Stain are compatible with mass spectrometry (MS) analysis.

To perform western blotting after InVision™ His-Tag In-Gel staining of His-tagged fusion proteins:

• Record a permanent image of the gel after staining of His-tagged fusion proteins.
• Equilibrate the gel in 1X SDS Running Buffer for 1 hour.
• Perform  western blotting and immunodetection using a method of choice.

E-PAGE™ gels are thicker than standard mini-gels and result in too much background when stained with InVision™ His-Tag In-Gel Stain. To obtain better staining sensitivity, we recommend transferring proteins of E-PAGE™ gels onto a nitrocellulose membrane and then staining the blot with the InVision™ His-tag In-gel Stain as described on page 14 in the manual.

We recommend storing the Lumio™ Green Detection kit components at –20 degrees C, protected from light, where it is stable for six months.

The Lumio™ Green Detection kit is a sensitive and highly specific kit for labeling Lumio™ fusion proteins prior to electrophoresis. It enables immediate visualization of Lumio™ fusion protein bands directly in a polyacrylamide gel.

The maximum excitation wavelength for Lumio™ Green Detection Reagent is at 500 nm and maximum emission wavelength is at 535 nm.

The color of the Lumio™ Green Detection Reagent may change from colorless to pink during storage. This color change does not affect the functioning of the reagent.

The kit is specially formulated for fast, sensitive, and specific detection, and is capable of detecting 1 pmole of a Lumio™ fusion protein (e.g., 1 pmole of a 30 kDa protein is 30 ng). The signal intensity may vary and is dependent on the individual protein. The signal intensity is also dependent on the number of moles of protein contained in a protein band because 1 molecule of Lumio™ Green Reagent binds to only 1 Lumio™ tag on the protein. For example, if you load 150 ng/band of 2 proteins with molecular weights of 150 kDa and 30 kDa, respectively, after detection with Lumio™ Green Detection Kit, the 30 kDa band fluoresces more intensely than the 150 kDa band. This is because there is only 1 pmole of the 150 kDa band while there are 5 pmoles of the 30 kDa band in the total mass loaded (150 ng/band).

The Lumio™ tag is small (6 amino acids, 585 Da). The small size of the tag is unlikely to interfere with the structure or biological activity of the protein of interest.

Proteins detected with the Lumio™ Green Detection Kit are compatible with mass spectrometry (MS) analysis. After detection with Lumio™ Green Detection Kit, excise the protein band/spot and prepare the samples for MS analysis using a method of choice or as directed by your core facility. The Lumio™ Green Detection protocol produces the following protein modifications. Be sure to account for these during MS analysis:

• Cysteines in the protein are modified and will result in the addition of 97.07 Da to each cysteine.
• The total molecular weight of the Lumio™ tag with the Lumio™ Green Reagent is 1060 Da (molecular weight of the Lumio™ Green Reagent without EDT is 475 Da and molecular weight of the Lumio™ tag is 585 Da).

To visualize Lumio™-tagged protein bands after staining, you will need a:

• UV transilluminator (302 nm) equipped with a standard video camera, CCD (Charged Couple Device) camera, or a cooled CCD camera with ethidium bromide filter or SYBR™ Green filter. Note: You can use a 365 nm UV transilluminator, but you may have to expose the gel for a longer time, as the sensitivity is lower than using 302 nm UV transillumination.

or

• Laser-based scanner with a laser line that falls within the excitation maxima of the stain (500 nm), and a 535 nm long pass filter or a band pass filter centered near the emission maxima of 535 nm. The sensitivity of detection is higher with laser-based scanners equipped with appropriate filters than with UV transillumination.

Samples prepared in standard (Laemmli) SDS-PAGE sample buffer are not compatible for use with the Lumio™ Green Detection Kit.

The Lumio™ In-Gel Detection Enhancer is a proprietary solution and is designed to reduce the non-specific binding of Lumio™ Green Detection Reagent with endogenous proteins.

The BenchMark™ Fluorescent Protein Standard allows direct visualization of molecular weight ranges of Lumio™ fusion proteins on an SDS-PAGE gel. The proteins in the standard are easily detected using a UV transilluminator or a laser-based scanner at the same excitation and emission wavelengths as your Lumio™ fusion protein.

The 97 Da molecular weight change is due to the Enhancer capping the cysteines that aren't part of the Lumio™ tag to prevent non-specific binding of the dye to the cysteines. So all cysteines except the 4 in the Lumio™ tag will be modified and show this change in molecular weight.

We recommend storing the Pierce™ 6xHis Protein Tag Stain Reagent Set at room temperature where it is stable for a year.

The Pierce™ 6xHis Protein Tag Stain Reagent Set can detect as low as 0.2 μg of a 35 kDa (5.7 pmol) His-tagged protein per band using a CCD camera, and as low as 2 μg (57 pmol) of the His-tagged protein per band with a UV transilluminator. Detection requires illumination of the stained gel with UV-light at a wavelength in the range 280–310 nm.

There is no fixation step in the Pierce™ 6xHis Protein Tag Stain Reagent Set staining procedure. Hence, staining with this kit does not inhibit subsequent total protein staining with general protein stains, or electrophoretic transfer to membrane.

Note: Bis-Tris gels run in MOPS or MES buffer may require fixing in 50% methanol: 7% acetic acid for 15 minutes before performing the stain procedure. After electrophoresis, fix the gel and then proceed with Step 1 of the procedure.

The fluorescent signal is stable for several hours in gels stored in water. Signal may be detectable, if somewhat attenuated, after overnight storage.

The gel would need to be sequentially stained and imaged after each stain. The order of staining would be:

InVision™ His-Tag In-Gel Stain (Cat. No. LC6030) → Image→ Pro-Q™ Diamond Phosphoprotein Gel Stain → Image → Pro-Q™ Emerald 300 or 488 Glycoprotein Gel Stain → SYPRO™ Ruby Protein Gel Stain → Coomassie™ Blue or Silver Stain → Image

The best step for leaving the gels overnight is during the fixation step, as the methanol and acetic acid both precipitate proteins and prevent diffusion. Gels are stable indefinitely in the fixation solution as long as the containers are well sealed to prevent contamination or gel drying and the containers are allowed to sit or rock gently to minimize gel damage. The high methanol concentration will dehydrate the gel, shrinking it and possibly giving it an opaque, white appearance. This is normal. Simply gently rock the gel in the wash solution to rehydrate to its original appearance.

Gels can also be left overnight in the water wash after the fixation step. After the post-fix wash, it is best to complete the staining procedure following the recommended protocol times. Once the gels are stained, the signal should be visible for at least several days, as long as the gels are protected from light. Stained and dried blots can be archived and the signal detected indefinitely, as long as the blots are protected from light.

No, the Pro-Q™ Diamond Phosphoprotein Gel Stain and Blot Stain are very different formulations and will not give acceptable phosphoprotein detection results on the alternate format.

Yes. We recommend staining with SYPRO™ Ruby Blot Stain after staining with Pro-Q™ Diamond Phosphoprotein Blot Stain.

Other known phosphoproteins can be used as positive control standards for the Pro-Q™ Diamond Phosphoprotein stain. Ovalbumin, in the Protein Molecular Weight Standards Reagent (Cat. No. P6649) is a phosphoprotein. None of the proteins in the Mark12™, Novex™ Sharp, SeeBlue™ or SeeBlue™ Plus2 standards is a phosphoprotein that could be used as a positive control with the Pro-Q™ Diamond Phosphoprotein stain.

The periodic acid oxidizing reagent breaks the bonds between vicinal hydroxyls (1,2 diols) on sugar residues via the formation of a cyclic periodate ester, forming either an aldehyde or ketone carbonyl group. The Pro-Q™ Emerald reagent contains a primary amine that will bind directly to the aldehyde/ketone, thus forming a covalent bond between the sugar residue and the dye molecule.

Note: The exact structure of the Pro-Q™ Emerald reagents is proprietary.

Pro-Q™ Emerald Glycoprotein gel stains work best with Tris-Glycine and Tricine gels. Pro-Q™ Emerald stains can be used with Novex™ NuPAGE™ Bis-Tris and Tris-Acetate gels, but may result in higher background staining, especially in combination with MES running buffer or in gels that are nearing their expiration date. If you wish to use Pro-Q™ Emerald stains with Novex™ NuPAGE™ Bis-Tris gels, we recommend using recently purchased gels and MOPS running buffer. The gel background increases with acrylamide density and gradient gels will show a gradual increase in background from the top to the bottom of the gel corresponding to the acrylamide gradient. Glycoproteins will still be detected in gels with high background, but with reduced sensitivity.

The gel would need to be sequentially stained and imaged after each stain. The order of staining would be:

InVision™ His-Tag In-Gel Stain (Cat. No. LC6030) → Image→ Pro-Q™ Diamond Phosphoprotein Gel Stain → Image → Pro-Q™ Emerald 300 or 488 Glycoprotein Gel Stain → SYPRO™ Ruby Protein Gel Stain → Coomassie™ Blue or Silver Stain → Image

Pro-Q™ Emerald 300/488 Glycoprotein Gel Stains are compatible with mass spectrometry. Only a minority of peptides in a trypsin digest will be glycosylated and the majority of non-glycosylated peptides can be identified. Glycosylated peptides are not detected under standard conditions (stained or unstained), as the databases do not take into account the glycans attached. One can deglycosylate the protein, which removes the covalently-bound dye and renders the peptide identifiable.

If the gel buffer contains Tris, glycine or any other component that has a primary amine in its structure, the primary amine will react directly with the aldehydes/ketones formed upon periodic acid oxidation, effectively capping all reactive sites and leaving no sites for the Pro-Q™ Emerald dye reagent to bind to.

Residual periodic acid must be removed from the gel prior to staining with Pro-Q™ Emerald 300/488 Glycoprotein stains so as to limit oxidation/decomposition of the Pro-Q™ Emerald dye.

SDS must be removed from the proteins prior to staining with Pro-Q™ Emerald 300/488 Glycoprotein stains so as to make the sugar residues accessible to the oxidizing reagent (periodic acid) and the Pro-Q™ Emerald dye reagent. SDS coats proteins and may limit access of the kit reagents by simple steric hindrance.

Other known glycoproteins can be used as positive control standards for the Pro-Q™ Emerald 300/488 Glycoprotein stains. Ovalbumin and transferrin in the Protein Molecular Weight Standards Reagent (Cat. No. P6649) are glycoproteins. None of the proteins in the Mark12™, Novex™ Sharp, SeeBlue™ or SeeBlue™ Plus2 standards is a glycoprotein that could be used as a positive control with the Pro-Q™ Emerald 300/488 Glycoprotein stains.

The best step for leaving the gels overnight is during the fixation step, as the methanol and acetic acid both precipitate proteins and prevent diffusion. Gels are stable indefinitely in the fixation solution as long as the containers are well sealed to prevent contamination or gel drying and the containers are allowed to sit or rock gently to minimize gel damage. The high methanol concentration will dehydrate the gel, shrinking it and possibly giving it an opaque, white appearance. This is normal. Simply gently rock the gel in the wash solution to rehydrate to its original appearance.

Gels can also be left overnight in the acetic acid wash after the fixation step. After the post-fix wash, it is best to complete the staining procedure following the recommended protocol times. Once the gels are stained, the signal should be visible for at least several days, as long as the gels are protected from light. Stained and dried blots can be archived and the signal detected indefinitely, as long as the blots are protected from light.

PVDF membranes must be used with Pro-Q™ Emerald 300/488 Glycoprotein stains. Nitrocellulose membranes will be dissolved by the high methanol in the fixation step.

Yes. We recommend staining with SYPRO™ Ruby Blot Stain after staining with Pro-Q™ Emerald 300 or 488 Glycoprotein Blot Stain.

The Thermo Scientific™ Pierce™ Power Stainer (Cat. No. 22833) consists of a Thermo Scientific™ Pierce™ Power Station (Cat. No. 22838) with activated Staining Software and a Thermo Scientific™ Pierce™ Power Stain Cassette (Cat. No. 22836). It is designed for rapid Coomassie staining of proteins in polyacrylamide gels and subsequent removal of stain from the gel background. Traditional Coomassie staining techniques require one hour to overnight of staining and destaining to achieve the desired results. When used in conjunction with Thermo Scientific™ Pierce™ Midi and Mini Gel Power Staining Kits, the Pierce™ Power Stainer is designed to provide staining efficiency in as few as 6 minutes that is equivalent to, or better than, traditional Coomassie™ staining techniques. This significant reduction in protein staining time is accomplished by fixing the protein to the gel and electrophoretically transporting the negatively charge Coomassie R250 dye rapidly through the gel matrix. The dye passes through the polyacrylamide and ionically binds to the protein, resulting in crisp blue bands with minimal background. The system has been verified to work with commonly used pre-cast and homemade SDS-PAGE gels.

The Pierce™ Power Stainer is designed to stain one to two mini-sized gels or one midi-sized gel in as few as 6.0 minutes. Please note that gels stained simultaneously must have the same formulation.

Yes. The Thermo Scientific™ Pierce™ Power Blot Cassette (Cat. No. 22835) can be purchased and will activate the pre-loaded Power Blotter software once inserted into your device.

Detection limits are similar to those of commercially available protein stains (~0.01 to 50 µg of pure protein).

The Pierce™ Power Stainer Cassette accommodates the staining for 1 or 2 Mini gels or 1 Midi gel.

Mini gels with dimensions of 7 x 8.4 mm and Midi gels with dimensions of 8 x 13.5 cm are compatible with Mini and Midi gel pads. Gel thickness of 1 mm is the most common. Gels with a thickness greater or less than 1 mm would require time optimization.

Pierce™ Power Staining Kits have enough material for 30 Mini gels or 15 Midi gels using the corresponding kit.

No. For the best results, use freshly prepared staining and destaining pads. Discard the used pads after each staining.

No. The Mini and Midi Gel Pads (8 layers per pad) are required to be used with the Pierce™ Power Stainer, Power Stain Solution and Destain Solution. The gel pads act as reservoirs for Power Stain Solution and Destain Solution. Standard western blotting filter paper will cause uneven staining and patchy background.

Coomassie R-250 dye is used. When optimized voltage is applied to the platinum-coated anode and stainless steel cathode, the negative charge of Coomassie R-250 dye causes it to travel through the gel and bind to the proteins. Unbound dye continues through and out of the gel matrix to destain the background.

No. The system will not work with regular Coomassie stain.

No. The Pierce™ Power Stainer requires the optimized and proprietary Power Stain Solution and Destain Solution.

Yes. Just place the gel in the Coomassie stain for 30–60 minutes and destain according to the manufacturer's protocol.

Gels stained using the Power™ Stainer can be stored in a plastic protective sheet or in water for up to 4 hours without loss of signal.

Yes. It is compatible with mass spectrometry with similar results to conventional Coomassie staining.

Yes. The Pierce™ Power Stain Cassette (Cat. No. 22836) can be purchased and will activate the pre-loaded Power Stainer software once inserted into your device.

No. During the staining, the proteins are fixed to the gel thus the western blotting results would not be optimal. The unstained gel should be used for western blotting.

The Thermo Scientific™ Pierce™ Power System (Cat. No. 22830) consists of the Pierce™ Power Station (Cat. No. 22838) with activated Staining and Blotting Software, the Pierce™ Power Stain Cassette (Cat. No. 22836), and the Pierce™ Power Blot Cassette (#22835). It is a combination of the Power Stainer and the Power Blotter.

Yes. The Pierce™ Power Station automatically recognizes the cassette that is inserted and initializes the correct software menu system.

No. It is not possible to insert the Power Blot Cassette and access the Power Staining protocol or vice versa.

No. Each cassette can only be used for its intended purpose; Power Blot Cassette for blotting (fast protein transfer) and Power Stain Cassette for protein staining. We have optimized the surface area to yield the best results for each application. The smaller stain cassette works better for staining while the larger blot cassette works better for blotting.

You can purchase the Thermo Scientific™ Pierce™ G2 Fast Blotter – Power Stainer Upgrade Kit (Cat. No. 62288). This kit contains the Pierce Power Stain Cassette and a USB Flash Drive loaded with software that will update the Pierce G2 Fast Blotter Control Unit software and add the staining functions. We also offer Thermo Scientific™ Pierce™ Power Staining Kits (cat. Nos. 22839, 22840).