MedSpace

Egg shell | Industrial applications of egg shell

noreply@blogger.com (MedSpace) — Mon, 18 Nov 2019 04:50:00 +0000

EGGSHELL CHARACTERISTICS:

Eggshell waste includes the shell and a membrane that separates the shell from the albumen (the white). These materials consist of as follows:

• 94% to 97% calcium carbonate;

• 3% to 6% protein;

• 1% various minerals (including magnesium, potassium, and traces of iron, sulfur and phosphorus).

Eggshell

APPLICATIONS AND POTENTIAL MARKETS FOR THE DIFFERENT EGGSHELL COMPONENTS:

Emerging sectors for eggshell application

Source:https://doi.org/10.1016/j.resconrec.2016.09.027

Stem cells promote peripheral nerve regeneration

noreply@blogger.com (Anonymous) — Fri, 11 Jan 2019 02:34:00 +0000

This is an embedded Microsoft Office presentation, powered by Office Online.

Patterning factors | determine regional identity and differentiation potential

noreply@blogger.com (Anonymous) — Mon, 19 Nov 2018 07:26:00 +0000

Ptf1a | Direct reprogramming of fibroblasts into neural stem cells

noreply@blogger.com (Anonymous) — Mon, 19 Nov 2018 07:21:00 +0000

RNA isolation from cells | RNA quantification

noreply@blogger.com (Anonymous) — Fri, 26 Oct 2018 06:59:00 +0000

RNA isolation from cells

Protocol:

Aspirate off media, wash once with ice cold PBS (1-2) mL.
Aspirate off PBS, add 1 mL Trizol.
Scrape the plate with a cell scraper, collect the cell lysate to 1.5 mL E-tube.
Incubate RT for 5mins.
Centrifugation 4°C, 13000 rpm for 10 mins.
Aliquot supernatant to new E-tube.
Add 200 µL chloroform (200 µL + 800 µL=1 mL), vortexing.
Incubate RT for 10-15 mins (vortex every 2 mins)
Centrifugation 4°C, 13000 rpm for 15 mins.
Separate aqueous phase and transfer to new E-tube.
Add isopropanol (equal volume of aqueous phase)
Inverting, incubate RT for 5-10 mins.
Centrifugation 4°C, 13000 rpm for 10 mins.
Remove supernatant and add 700 µL 70%EtOH to wash the pellet.
Centrifugation 4°C, 13000 rpm for 5 mins.
Remove supernatant and add 1 mL 70%EtOH to wash the pellet.
Centrifugation 4°C, 7000 rpm for 10 mins.
Remove supernatant, spin down.
Use yellow tip remove supernatant completely.
Air dry rapping with poly for 5-10 mins.
Add DEPC-DW 35 µL to E-tube.
Spin down, qty/store -80°C.

Fig: Illustration of RNA isolation from cells

RNA quantification

Instruments:

20 µL pipet
100 µL pipet
Yellow & white tip
Tip discard box
Kim tissue
70% EtOH
75 µL tube & levelling
RNA sample
DEPC-DW

Fig: Denovix DS-11 Spectrophotometer from DeNovix Inc.

Protocol:

Place DEPC-DW & RNA sample on ice.
Turn on NANODROP machine (Location: 702)
Open and enter RNA button
Wipe the microplate using 70% EtOH
Add, 2 µL DEPC-DW to the microplate, press the measure button, then wipe the plate with 70% EtOH.
Touch the BLANK, wipe.
Add, 2 µL RNA sample to the microplate, press the measure button, then wipe the plate with 70% EtOH.
Store and transfer data to USB.
Calculate the vol. Of RNA samples for reverse transcription in excel.

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Western blot | troubleshooting | samples turn yellow and precipitate

noreply@blogger.com (Anonymous) — Wed, 18 Jul 2018 08:07:00 +0000

Why western Blot samples turn yellow and precipitate?

Sample appears to be acidic. Bromophenol blue in the loading buffer turns to yellow at acidic pH (below 3). Try to adjust the pH by adding NaOH [1].
Reason for that is proteins may be getting degraded due to heat exposure. One researcher had the same problem and so instead of keeping samples at 95°c he started warming at 60-65° C. So anyone can try it out and as [1] said check the pH as well of the loading dye [2].
Heating a sample also changes pH. An altered pH is not only responsible to turn the sample color yellow, but may also precipitate the proteins. Keeping SDS containing samples in ice is another common mistake to see this precipitation [3].
Another researcher said that he has got to do more with the total protein concentration in the lysate. He faced the same problem of precipitation upon denaturation at 100°c. He added 1/10th volume of 10% SDS and then boiled. Protein did not precipitate. The high concentration of SDS will solubilize the proteins and prevent their precipitation [4].

Western blot | Re-use a primary antibody

noreply@blogger.com (Anonymous) — Wed, 18 Jul 2018 07:18:00 +0000

Why reuse primary antibody?

Western blotting is one of the most common method for detecting and semi-quantifying specific proteins. The cost of running a Western is predominantly driven by detection antibodies, which are used to probe membrane-bound target protein(s). Although some antibodies can be produced cheaply in-house, most antibodies must be commercially ordered. Reusing antibodies for subsequent Westerns can save your lab a considerable amount of money. So money is nothing but everything. In conclusion economic research practises is budget friendly and reuse of primary antibody is one of them.

When reuse is problematic?

The most standard phospho-antibody protocols involve blocking with 5% milk in PBS or TBS (0.1% Tween 20), followed by a couple washing steps with PBS or TBS to remove the excess milk. If you dilute your antibody in a milk solution, storage and re-use will be extremely problematic, because the milk will "go bad" and fall out of solution, similar to milk at home.
BSA is ideal for your needs. Dilute your primary antibody in 5% BSA in PBS or TBA (0.1% Tween 20) [1].
Generally milk is not used for blocking if its a phospho antibody. You can store used antibody in 4C for upto 2 weeks [2].

When and how we reuse primary antibody?

Use 1% non-fat milk for antibody preparation. If you want to use further then give pri Ab treatment at 4 deg for overnight or at least 4 h. Later, store at -20 deg. You can use easily 2-3 times. It works for most of the antibodies [3].
Both myself and my lab mates dilute our primary antibodies in 5% BSA in TBST with Sodium Azide stored at 4C for months to over a year and typically use them several times a month to weekly with great results [4].
Hii, Dilute primary antibodies in TBST with 5% BSA. Usage of milk can create non-specific background, because casein itself is a phosphoprotein. Further, milk in TBST can not be stored for long at 4 degree C. you can store your primary antibody at 4 degree C for 2-3 weeks and can reuse 2-4 times efficiently [5].
You may use milk but BSA is better if you plan to store your diluted antibody longer. Use sodium azide (0.05%). Even with azide, milk in your dilution buffer will get spoiled thereby affecting long term storage. Have you ever tried diluting your phospho antibodies in just plain PBS without any blocking agent? You are probably doing the blocking step prior to your antibody step right?[6].
It all depends on how strong the signal you get with the previously used antibodies and how stable they are. Also, it depends on the working titer of the antibodies and how much of the antibodies were consumed during your first use of them. My advice to everybody is to standardize that by rechecking the reactivity of previously used antibodies on strips from the same target protein and see if the signal remains as strong or gets weaker upon use [7].
We design and produce phospho antibodies in our lab. We block our antibodies with serine and threonine phospho sites with 5% non-fat milk. We use BSA for phospho tyrosine antibodies because the phospho tyrosine in milk can produce high background. It is true that milk goes bad faster than BSA, and BSA will certainly work for all antibodies. The azide is great as long as it doesn't interfere with your system. We divide our antibodies into working aliquots, and only keep one aliquot in 4 degrees at a time. As mentioned in other posts, we add glycerol to 50% and store at -20 to avoid freeze/thaw. For long term storage, we recommend aliquoting and storing at -80. We generally don't reuse our antibodies, but here is an idea to reduce the amount of antibody you use when testing multiple antibodies using the same lysate: Try a gel with a stacking layer that has one large trough instead of one that has multiple lanes. Once the gel has been transferred to a blot, you can cut the blot into strips of equal widths. In most cases, twice as many strips can be cut from one large trough than from cutting apart a blot with multiple lanes. The strips are also more narrow and require much less incubation solution [8].

Western blot | Bradford assay ⎸Protein quantification

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:58:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Bradford assay ⎸Protein quantification

A. Preparation of BSA standard curve

Protocol:

Preparation of 10 mg/ml BSA solution (10 ml DW + 100 mg BSA) [Stock solution]
Prepare 1 mg/ml BSA solution (9 ml DW + 1 ml of 10 mg/ml BSA) [Working solution]
Levelling total 8 cuvettes as Blank, 1, 2, 4, 6, 8, 10, 12 µL respectively.
For blank, add 1 mL lysis buffer. And, add 1 mL working solution to 1, 2, 4, 6, 8, 10, 12 µL levelled cuvettes respectively.
Discard 1, 2, 4, 6, 8, 10, 12 µL working solution from 1, 2, 4, 6, 8, 10, 12 µL levelled cuvettes gradually.
Start quantification and get a BSA curve.

B. Protein quantification

Protocol:

Part-1 (Prepare working solution):

Prepare lysis buffer (Western blot protocol)
Mix the tissue with lysis buffer in a conical tube (10 mg tissue/1 mL lysis buffer) and incubate for one hour.
Prepare bradford working solution by mixing bradford stock solution and autoclaved DW with a ratio of 1:4 [{Dilution formula, M1V1 = M2V2}; {200µL ✖ 1mL = 800µL ✖ 1mL}] [Working solution]
Centrifuge at 13000 rpm in 4℃ for 20 mins
Transfer the supernatant to new e-tube

Part-2 (Start quantification):

Aliquot 1 mL (100 µL) of bradford working solution.
Discard 1 µL of bradford working solution
Add 1 µL of sample [that you want to quantify]
Gently pipetting and incubate for 5 mins
Set the wavelength of spectrophotometer 595 nm
Quantification.

Western blot | ECL development

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:54:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

ECL development

Protocol

Remove ECL Western Blotting Reagent Kit (Pierce® ECL plus western blotting substrate) from the refrigerator.
Prepare the substrate working solution by mixing Substrate A and Substrate B in a 40:1 ratio (example: 12 mL substrate A + 300 µL substrate B for 8 × 12 cm2 membrane). Use 0.125mL working solution per cm2 of membrane.

Note: The working solution is stable for up to 1 hour at RT.

Place the membrane on the other suitable clean surface, remove the water, add Pierce® ECL plus western blotting substrate (working solution) and spread evenly over the membrane surface.
Incubate blot with working solution for 5 minutes at RT.
Remove blot from working solution and place it in a plastic sheet protector or clear plastic wrap. Use an absorbent tissue to remove excess liquid and carefully press out any bubbles from between the blot and the membrane protector.
Place the protected membrane in a film cassette with the protein side facing up. Turn off all lights except those appropriate for X-ray film exposure (e.g., a red safelight).

Note: Film must remain dry during exposure. For optimal results, perform the following precautions:

• Make sure excess substrate is removed from the membrane and the membrane protector.
• Use gloves during the entire film-handling process.
• Never place a blot on developed film, as chemicals on the film may reduce signal.

Cut the X-ray film up to the membrane size (fold the top of the X-ray film and mark the direction).
Carefully place X-ray film on top of the membrane. Perform a first-time exposure of 60 seconds. Vary the exposure time to achieve optimal results.

Note: Light emission is most intense during the first 5-30 minutes after substrate incubation. Light emission continues for several hours but decreases with time. Longer exposure times may be necessary as time elapses.

Caution: Any movement between the film and membrane can cause artifacts on the film.

Develop film using appropriate developing solution and fixative. If the signal is too intense, reduce exposure time or strip and reprobe the blot with decreased antibody concentrations.
Rinse the X-ray film thoroughly with flowing water and air-dry. Be careful that the water does not stick together.

Western blot | Antibody incubation & Washing

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:51:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Antibody incubation & Washing

※ When you first take the Antibody, it tells you how much concentration should be diluted. Use it at that concentration. If the protein band comes out, continue to use it. If the band does not come out, increase the concentration.

Primary & secondary antibody incubation

※ If there are several proteins you want to see, the membrane may be cut to size. Prior to Primary Ab treatment, remove the water, wrap it in a wrap, and check for protein markers (total volume 2-3 ml).

A. Dilution of primary Ab:

Dilute primary Ab in TBST (0.5% Skim milk / TBST) and incubate shaking incubation for 1 hour at room temperature. Or overnight at 4 ° C.

→ 0.5% skim milk + TBST + primary Ab

→ Used primary Ab can be re-used, can be kept at 4 ℃

Ex 1) TBST 10 ml + Skim milk 0.05 (g) + 10 μl of primary AB when the dilution concentration is 1: 1000 and the total volume is 10 ml

Ex 2) TBST 2 ml + Skim milk 0.01 (g) + 2 ㎕ of primary AB when the dilution concentration is 1: 1000 and the total volume is 2 ml.

➢ The membrane is washed three times for 10 minutes with TBST buffer

→ If the membrane is turned upside down, it will not wash properly.

→ If the membrane is cut, use 3-5 ml

B. Dilution of Secondary AB:

dilute the secondary AB in TBST (or 1% skim milk / TBST), then shake the membrane at room temperature for 1 hour.

→ TBST + primary Ab

Ex 1) When the dilution concentration is 1: 1000 and the total volume is 10 ml, TBST 10 ml + 10 ㎕ of secondary Ab

Ex 2) When the dilution concentration is 1: 1000 and total volume is 2 ml, TBST 2 ml + 2nd Ab 2 ul.

2.Washing:

➢ The membrane wash 5 times with TBST solution every 10 minutes.

→ If the membrane is inverted, the washing will not work properly

C. Washing:

➢ The membrane wash 5 times with TBST solution every 10 minutes.

→ If the membrane is inverted, the washing will not work properly.

Western blot | Protein transfer

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:47:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Transfer membrane

Preparation:

• 10X & 1X Transfer buffer (4℃)

• Sponge / Filter paper / 3MM paper / Membrane

• Cassette / Blotting chamber

• 10X TBS (pH 7.5) & 1X TBST

• 0.2% Ponceau S

• Coomassie blue

• Blocking buffer (5% skim milk in TBST)

• Staining solution & Destaining solution

• BM purple AP substrate

Transfer protocol:

Cut the membranes to the appropriate size and immerse them in transfer buffer.
For PVDF membrane, immerse in methanol for 5 minutes before transferring to transfer buffer.

Membrane activation: The process by which proteins in the gel make hydrophobic interaction with the membrane. This process is not necessary when using an NC.

Transfer the electrophoresis gel to the transfer buffer, remove the wells, and remove the remaining portion except the sample. → Save membrane.
Place the black portion of the Western blot cassette against the bottom of the container containing the transfer buffer and place it on top of it:

→ Immersed in transfer buffer accordingly, dacron sponge / filter paper (1 sheet) / 3MM paper (2 sheets) / gel / Membrane / 3MM paper (2 sheets) / filter paper (1 sheet)/dacron sponge.

→ The gel of the black part of the cassette is the cathode and the membrane of the white part is the anode.

5. Carefully raise the gel on 3MM paper to avoid air bubbles. At this time, I think about the direction of the transfer. If you move the marker to the left, transfer it later and do not reverse the direction when reading the result. → Push the spreader slightly so that the bubble disappears.

6. Carefully raise the membrane on the gel to prevent air bubbles. Place 2 sheets of 3MM paper, 1 filter paper and dacron sponge on it, and gently push it with a spreader to release the bubble.

7. Carefully fasten the cassette to prevent air bubbles from forming and then tighten the clip so that both cassettes do not fall off.

8. After transferring the cassette to the blotting chamber, insert the 1X transfer buffer so that the membrane in the cassette is sufficiently immersed. Place the electrode so that the membrane is the anode and the gel is the cathode.

9. Fill the Bio-Ice cooling unit with ice to prevent denaturation of protein due to heat generated during transfer.

10. Transfer with 230 mA for 1 ~ 1.5 hr.

→ The larger the desired protein size, the longer the time required for the transfer.

→ Prepare the blocking buffer 10 minutes before the transfer is completed.

11. After transfer, remove the membrane, rinse briefly in TBST, and incubate in a compact racker containing blocking buffer (5% skim milk in TBST) for 1 hour at room temperature.

→ It reduces the background by blocking the unbound portion of the membrane in the membrane.

→ If there is no time, put in blocking buffer and overnight at 4 ℃.

→ 2 gels require blocking buffer 30 ml, skim milk 1.5 g (5%) and BSA (-20 ° C storage) 300 μl (1%)

TBST 30 ml + Skim milk 1.5 g → Vortexing + 1% BSA (300 ul)

12. After the transfer, remove the membrane and immerse it in 0.2% Ponceau S to stain.

→Mark the marker area with a marker pen on the dyed marker area.

13. Coomassie staining of transferred gel is used to analyze the cause of failure.

Sandwich assembly:

+ve electrode

Sponge

Filter paper

3MM paper

PVDF membrane

Gel

3MM paper

Filter paper

Sponge

-ve electrode

Western blot | Ponceau S Staining

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:44:00 +0000

Lab. of Bioclock & Aging/Dewan Md. Sumsuzzman

Ponceau S Staining

Ponceau S Staining Solution:

Table-1: Recipe of 0.1% (w/v) Ponceau S in 5% (v/v) Acetic acid

Ingredients	Quantity
Ponceau S	1 gm
Acetic acid	50 ml
ddH2O	Make up to 1L

Table-2: Recipe of 0.2% (w/v) Ponceau S in 3% (v/v) Acetic acid

Ingredients	Quantity
Ponceau S	2 gm
Acetic acid	30 ml
ddH2O	Make up to 1L

Table-3: Recipe of 0.1N NaOH in 100 ml water

Ingredients	Quantity
NaOH	0.4 gm
H2O	100 ml

Protocol:

Following Western blotting, transfer the membrane to 5ml Ponceau S Stain solution.
Place on an orbital shaker for 5 minutes at room temperature.
Rinse membrane with DI water to achieve desired staining, approximately 2‐3 washes of 5 minutes each will remove the background staining.
For total protein destaining, use a 0.1N NaOH solution. Wash the membrane with 0.1N NaOH solution for 5 minutes. Protein bands will start disappear after 10-30 seconds. Discard wash solution and repeat once.
Wash the membrane 2‐3 times with DI water for 5 minutes each.
Continue with Western blot/ immunoblot procedure.

Note:

The product can be used on most membranes including PVDF (polyvinylidene difluoride) except nylon membranes. Ponceau S does not destain from nylon membranes because the nylon is positively charged, the stain is negatively charged, and therefore, the stain is electrostatically held. Other membranes, like nitrocellulose, cellulose acetate, or PVDF are electrically neutral, and therefore, can easily be destained.

A quick water rinse allows one to see if bands are developing, but destaining is usually done in 10% acetic acid. For sequencing or immunoassay purposes, the bands can be destained to colorless using 0.1 M NaOH.

If using nitrocellulose membranes, do not use alcohols in destaining procedure. For a western blot, rinsing the membrane with deionized water will remove any background dye. To remove the dye from the protein for further immunochemical testing, a rinse the membrane with 0.1 M NaOH. Destaining in acetic acid is not done on the nitrocellulose membrane.

Western blot | Coomassie Blue Staining

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:39:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Coomassie Blue Staining

Table-1: Recipe of Staining Solution

Ingredients	Quantity
Coomassie Blue R-250	2.5 gm
Methanol	500 ml
Acetic acid	100 ml
DW	400 ml

Note: Final volume 1L.

Add 100 ml of acetic acid to 500 ml of ddH2O.
Add 500 ml of methanol and mix.
Add 1g of Coomassie R250 dye and mix.
Put a magnetic bar in a brown bottle, and add dye while spinning.
Finally, filter it.

Store at room temperature in a sealable container.

Table-2: Recipe of Destaining Solution

Ingredients	Quantity
Methanol	500 ml
Acetic acid	100 ml
DW	400 ml

Store at room temperature in a sealable container.

Protocol

After the gel has finished running, remove it from the tank and remove the gel from the glass plates.
Wash the gel 2-3 times, 5 minutes each in deionized water to remove SDS present in the gel. Each wash should be large volumes of water.
Remove all free water from the gel. Add an adequate amount of staining solution (Coomassie Blue R-250) to cover the gel. And put a lid on the container to prevent the Coomassie from evaporation.
Place the container on a rocking device for gently shaking the gel in staining solution for 1 hour. Protein bands will be visible within 3-5 minutes and reach a maximum intensity within 1 hour. Longer incubation may be performed.
Rinse the stained gel in a large volume of deionized water, 2-3 times for 5 minutes each. To de-stain, cover the gel in destaining solution and place on a rocking device for 30 minutes or until desired resolution is attained.
Rinse the gel in deionized water and image/store the gel.

Note: If background staining is noticed, it is indicative of residual SDS in the gel. Rinsing the gel extensively in deionized water will remove the background staining.

Western blot | Gel Electrophoresis

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:30:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Gel Electrophoresis

Preparation of Gel:

Part-1:

Clean glass plates with D.W and 70% ETOH, dry thoroughly and assemble them.
Place 100% ETOH between glass plates to check for leakage.
If the ETOH does not leak, clean the ETOH using 3MM paper.
Carefully insert the comb, mark it in the comb down to 0.5 cm, and remove the comb.

Part-2:

Identify the molecular weight of the protein to be separated and determine the concentration of acrylamide.

Table-1: Acrylamide concentration as per protein size.

Acrylamide concentration (%)	Protein size (kDa)
15	10-43
12	12-60
10	20-80
7.5	36-94
5	57-212

Table-2: Preparation of running gel.

Running Gel (10 ml)	5％	7.5％	10％	12％	15％
D.W H2O	5.3 mL	4.6 mL	4 mL	3.3 mL	2.3 mL
1.5 M Tris-Cl (pH 8.8)	2.5 mL	2.5 mL	2.5 mL	2.5 mL	2.5 mL
30 acrylamide Solution	2 mL	2.7 mL	3.3 mL	4 mL	5 mL
10 SDS	100 µL	100 µL	100 µL	100 µL	100 µL
TEMED	8 µL	6 µL	4 µL	4 µL	4 µL
10 APS	100 µL	100 µL	100 µL	100 µL	100 µL

Note: Before adding APS and TEMED to the solution, thoroughly mix the solution, settle it down for 5-10 mins and after then add APS and TEMED and mix the solution very gently to avoid any air bubble.

Running Gel is poured to the bottom of the comb to a depth of about 0.5 cm.
Carefully put 1 ml of Isopropanol (or 1 ml of water-saturated N-butanol) and let the gel dry.
After 30-40 minutes, remove the Isopropanol and wash with D.W.
After wiping with 3MM paper, re-mark the position of the running gel (let gel down a lot).

Table-3: Preparation of stacking gel.

5% Stacking gel	2ml	4ml	6ml	8ml
D.W H2O	1.4 mL	2.7 mL	4.1 mL	5.5 mL
1.5 M Tris-Cl (pH 6.8)	250 µL	500 µL	750 µL	1000 µL
30 acrylamide Solution	330 µL	670 µL	1000 µL	1300 µL
10 SDS	20 µL	40 µL	60 µL	80 µL
TEMED	2 µL	4 µL	6 µL	8 µL
10 APS	20 µL	40 µL	60 µL	80 µL

Note: Before adding APS and TEMED to the solution, thoroughly mix the solution, settle it down for 5-10 mins and after then add APS and TEMED and mix the solution very gently to avoid any air bubble.

Pour 5% stacking gel and carefully insert the comb so the bubble does not get in.
If you combine a stacking gel in a comb, you will get less bubble.
After inserting the comb, fill the stacking gel in the space where the stacking gel is not enough so that the well does not collapse.
After 30-60 minutes, if the stacking gel hardens, remove the comb carefully and clean the wells with a syringe to make it easier to load the protein into the wells.
Transfer the small glass into the electrophoresis tank with the inside facing, then fill the 1X SDS running buffer so that it does not leak.

B. Sample preparation & loading:

While the gel is firm, pre-quantify the protein sample (stored at -80 ° C) in ice and dissolve.
Set the temperature of the dri-bath to 95 ° C in advance.
Mix the same amount of 2X sample buffer with the protein sample.
Heating with above samples at 95 ° C for 10 min [Protein denaturation]
Cooling at ice (4 ℃) for 10 min.
Spin-down lightly, then load the gel and store the remaining sample at -80 ° C.

Table-4: Special recommendation for sample preparation.

Ingredients	Quantity	Recommendation
Marker	3-4 microL	Do not vortexing
Sample+2x Sample buffer	10 ul + 10 ul	Pipetting
Loading dye	10 ul (the same amount as the sample)	Heat block with 95 ° C for 10 min

C. Running the gel:

Stacking gel: 80-100 V (mainly 80 V to 20 min)
Running gel: 120V (about 1.5 hr) → run until the dye reaches the bottom of the gel, usually 20 minutes after running out of dye.

Western blot | Buffer preparation

noreply@blogger.com (Anonymous) — Thu, 28 Jun 2018 01:15:00 +0000

Lab. Bioclock & Aging/Dewan Md. Sumsuzzman

Buffer preparation

1. Lysis buffer-cell/tissue

Step 1: Preparation of 1 M of Tris-HCl (pH 7.4) stock solution

Dissolve 121.1 g Tris-base in 800 ml distilled water, adjust the pH to 7.4 using HCl solution, and make up the volume to 1000 mL using distilled water.
Autoclave to sterilize.

Step 2: Preparation of 100 mL 1M NaCl

Dissolve 5.844 g NaCl in 80 mL of H2O. and make up the volume to 100 mL using distilled water.

Step 3: Preparation of Sodium Deoxycholate, NaDc (10％, 50ml)

Dissolve 5 g Sodium Deoxycholate in 25 mL D.W. and make up the volume to 50 mL. Filtration and store.

Step 4: Preparation of 200 mM Sodium orthovanadate, Na3VO4 (200 mM, 50 mL)

Dissolve Sodium orthovanadate 1.84 gram in 45 mL D.W.

pH 10 (yellow) and boiling (until colorless), 10 min
cool to R / T
pH 10, Boiling for 10 min
cool to R / T and pH 10
finally, make up the volume to 50 ml,
aliquot by 1ml and store at -20 ℃

Step 5: Preparation of Sodium Fluoride, NaF ( 100mM, 50ml)

Dissolve 0.21 gram Sodium Fluoride (MW: 41.99) 0.21 gram in 25 mL of D.W.
finally, make up the volume to 50 ml,
Filtration, and
Storage in room temperature.

Step 5: Preparation of Protease inhibitor cocktail, PIC, (×7)

Dissolve Protease inhibitor cocktail tablet 1 in 1.5 ml D.W.
Vortexing, aliquot stored at -20 ° C

Step 5: Preparation of 10% NP-40

Mix 10 ml of NP-40 with 90 ml of ddH2O by stirring.
Store at 4°C.

Table-1: Lysis buffer-cell

SL/No.	Store	Content	Stock	1 mL	2 mL	4 mL
1	RT	50 mM Tris-HCL (pH7.4)	1 M	50 µL	100 µL	200 µL
2	4	1% NP-40	10%	100 µL	200 µL	400 µL
3	4	0.25% Sodium Deoxycholate, NaDc	1%	250 µL	500 µL	1000µL
4	RT	150 mM NaCl	1 M	150 µL	300 µL	600 µL
5	-30	1 mM Na 3 VO 4 (Sodium orthovanadate)	200 mM	5 µL	10 µL	20 µL
6	RT	1 mM NaF (Sodium fluoride)	100 mM	10 µL	20 µL	40 µL
7	-30	1-2 mM protease inhibitor cocktail	× 7	143 µL	286 µL	571 µL
8	RT	D.W.		292 µL	584 µL	1169 µL

Table-2: Lysis buffer-tissue

Sl/No.	Store	Content	Stock	10 mL
1	RT	50 mM Tris-HCL (pH7.4)	1 M (next, 100mM)	500 µL
2	RT	150 mM NaCl	1M (next, 300mM)	1.5 mL
3	RT	0.1% SDS	10%	100 µL
4	4	1% NP-40 (light protected)	10% in water (boiled 1ml/9ml)	1 mL
5	4	0.5% Sodium Deoxycholate, NaDc	10% in water (0.1 gram/1 mL)	500 µL
6	-20	1 mM Na 3 VO 4 (Sodium orthovanadate)	200 mM	50 µL
7	-20	5 mM NaF (Sodium fluoride)	500 mM (0.021 gram/1 mL)	100 µL
8	-20	protease inhibitor cocktail		1 tablet
9	RT	1 mM PMSF	200 mM in isopropanol (0.03484 gram/1 mL)	50 µL
10	RT	Autoclaved D.W.		6.2 mL

Note: PMSF use after 30 mins to dilution into buffer

For clarifying stock solution: Click here
For making working solution from stock: Click here

2. Buffer system

A. 30％ Acrylamide solution

Content	Conc.	50 mL	100 mL	200 mL
Acrylamide	29%	14.5 g	29 g	58 g
Bis-acrylamide	1%	0.5 g	1 g	2 g
Total	30%	15 g	30 g	60 g

Protocol:

It dissolves by applying heat.
Preparation: beaker, filter paper, bottle
Fill the beaker with foil and dissolve enough.
Filtration with Whatman No.1 filter (0.45 µm)
Put it in a brown bottle or a bottle filled with foil and
store at 4 ℃ (1 month or less)

B. 1 M Tris-Cl (Lysis buffer pH 7.4 / Stacking gel pH 6.8)

Autoclaved
Molecular weight of Tris-Cl (MW: 121.14),

121.14 (g) x 1 M = 121.14 (g)

1000 mL: 121.14 = 200 mL : X

X= 121.14 ✖ 200/1000

X= 24.228 g

S0, 24.228 g of Tris-Cl dissolve in 180 mL of D.W. Adjust pH with HCl. Make the volume up to 200 mL.

C. 1.5 M Tris-Cl (running gel pH 8.8)

Autoclaved
Molecular weight of Tris-Cl (MW: 121.14),

121.14 (g) x 1.5 M = 181.71 (g)

1000 mL: 181.71 = 200 mL : X

X= 181.71 ✖ 200/1000

X= 36.342 g

S0, 36.342 g of Tris-Cl dissolve in 180 mL of D.W. Adjust pH with HCl. Make the volume up to 200 mL.

D. 10% SDS

If the total volume is 200 ml, add 20 (g). Ex) 200 ml × 10 (%) = 20 (g)

E. 10X SDS running buffer (pH 8.3)

Content	10X	1X
250 mM Tris-base	30.28 g	3.028 g
1.92 M glycine	144.13 g	14.41 g
1％ SDS	10 g	1 g
Total volume	1 L	1 L
Adjust the pH to 8.3 with HCl and adjust the total volume to 1L.

To create 1X SDS running buffer immediately, calculate Con. of each buffer by 1/10.

25mM Tris base
192mM glycine
0.1％ SDS

F. 2× sample loading buffer (pH 6.8)

Content	1X	1X 10 mL	2X	2X 10 mL	20 mL
1M Tris-Cl (pH 6.8)	50 mM	1 mL	100 mM (0.1 M)	1 mL	2 mL
100％ Glycerol	10％	1 mL	20%	2 mL	4 mL
10％ SDS	2％	200 µL	4%	4 mL	8 mL
β-Mercaptoethanol	2.5％	250 µL	5%	500 µL	1 mL
4％ Bromophenol blue	0.1％	10 µL	0.2%	20 µL	1 mL
D.W.		7 mL 540 µL		2 mL 480 µL	4 mL

Immediately prior to use, add β-Mercaptoethanol.
Addition of 50 ul for 2X (because it is aliquoted by 1ml), addition of 25
ul for 4X

G. 10× Transfer buffer (pH 8.3), 1L

Content	Qty.
Tris (250 mM)	30.28 g
Glycine (1920 mM)	140 g

Final volume 1L, then store at 4 ℃.

H. 1× Transfer buffer (pH 8.3), 1L

Content	Qty.
10X transfer buffer	100 mL
MetOH	200 mL
D.W.	700 mL

MetOH must be added

I. 10× TBS (pH 7.5), 1L

Content	Qty.
NaCl (1.5 M)	87.66 g
Tris (100 mM)	12.114 g

Finalize total volume 1L, then store by autoclave at 4 ℃.

J. 1× TBST, 1L

Content	Qty.
10X TBS	100 mL
D.W.	900 mL
Tween-20	1 mL

H. PBS (Phosphate buffer saline, pH7.4)

Content	Qty.
Working conc.	800 mL
137 mM NaCl	6.4 g
2.7 mM KCl	0.16 g
10 mM Na 2 HPO	1.152 g
2 mM KH 2 PO 4	0.192 g
D.W. up to 800 ml

I. Blocking buffer (5% skim milk in TBST)

Two gels need 30 ml.
When the blocking buffer is 30 ml, BSA (stored at -20 ° C) enters 300 μl

Add BSA 1% along with desired quantity of TBST & skim milk.

TBST 500 mL + Skim milk 25 g → Vortexing

TBST 30 mL + Skim milk 1.5 g → Vortexing

TBST 20 mL + Skim milk 1 g → Vortexing

Bradford Protein Assay for determining protein concentration

noreply@blogger.com (Anonymous) — Thu, 21 Jun 2018 10:59:00 +0000

Principle:

The Bradford protein assay is used to measure the concentration of total protein in a sample. The principle of this assay is that the binding of protein molecules to Coomassie dye under acidic conditions results in a color change from brown to blue. This method actually measures the presence of the basic amino acid residues, arginine, lysine and histidine, which contributes to formation of the protein-dye complex. Unlike the BCA assay, reducing agents (i.e. DTT and beta—mercaptoethanol) and metal chelators (i.e. EDTA, EGTA) at low concentration do not cause interference. However, the presence of SDS even at low concentrations can interfere with protein-dye binding. 

Materials and Reagents

Bovine Serum Abumin (BSA) (Sigma-Aldrich)
Coomassie Brilliant Blue G-250 (Sigma-Aldrich, catalog number: 27815 )
Methanol
Phosphoric acid (H3PO4)
Bradford reagent (see Recipes)

Equipment

Spectrophotometer (Tecan)
Whatman #1 paper (Whatman)

Procedure

Standard assay procedure (for sample with 5-100 µg ml-1 protein)
1. Prepare five to eight dilutions of a protein (usually BSA) standard with a range of 5 to 100 µg protein.
2. Dilute unknown protein samples to obtain 5-100 µg protein/30 µl.
3. Add 30 µl each of standard solution or unknown protein sample to an appropriately labeled test tube.
4. Set two blank tubes. For the standard curve, add 30 µl H2O instead of the standard solution. For the unknown protein samples, add 30 µl protein preparation buffer instead. Protein solutions are normally assayed in duplicate or triplicate.
5. Add 1.5 ml of Bradford reagent to each tube and mix well.
6. Incubate at room temperature (RT) for at least 5 min. Absorbance will increase over time; samples should incubate at RT for no more than 1 h.
7. Measure absorbance at 595 nm.
Microassay procedure (<50 µg ml-1 protein):
1. Prepare five standard solutions (1 ml each) containing 0, 10, 20, 30, 40 and 50 µg ml-1 BSA
2. Pipet 800 μl of each standard and sample solution (containing for <50 µg ml-1 protein) into a clean, dry test tube. Protein solutions are normally assayed in duplicate or triplicate.
3. Add 200 μl of dye reagent concentrate to each tube and vortex.
4. Follow the procedure described above for the standard assay procedure.

Recipes

Bradford reagent Dissolve 50 mg of Coomassie Brilliant Blue G-250 in 50 ml of methanol and add 100 ml 85% (w/v) phosphoric acid (H3PO4). Add the acid solution mixture slowly into 850 ml of H2O and let the dye dissolve completely (note: Do not add H2O into the acid solution). Filter using Whatman #1 paper to remove the precipitates just before use. Store in a dark bottle at 4 °C.

Read more related articles MedSpace

References: http://www.bio-protocol.org

Cell Culture Environment

noreply@blogger.com (Anonymous) — Thu, 21 Jun 2018 10:48:00 +0000

Introduction

One of the major advantages of cell culture is the ability to manipulate the physicochemical (i.e., temperature, pH, osmotic pressure, O2 and CO2 tension) and the physiological environment (i.e., hormone and nutrient concentrations) in which the cells propagate. With the exception of temperature, the culture environment is controlled by the growth media. 

While the physiological environment of the culture is not as well defined as its physicochemical environment, a better understanding of the components of serum, the identification of the growth factors necessary for proliferation, and a better appreciation of the microenvironment of cells in culture (i.e., cell-cell interactions, diffusion of gases, interactions with the matrix) now allow the culture of certain cell lines in serum-free media. 

Adherent vs Suspension Culture:

A. Adherent Culture:

Appropriate for most cell types, including primary cultures. 

Requires periodic passaging, but allows easy visual inspection under inverted microscope. 

Cells are dissociated enzymatically (e.g., TrypLE™ Express, trypsin) or mechanically .

Growth is limited by surface area, which may limit product yields. 

Requires tissue-culture treated vessel. 

Used for cytology, harvesting products continuously, and many research applications. 

B. Suspension Culture:

Appropriate for cells adapted to suspension culture and a few other cell lines that are nonadhesive (e.g., hematopoietic). 

Easier to passage, but requires daily cell counts and viability determination to follow growth patterns; culture can be diluted to stimulate growth.

Does not require enzymatic or mechanical dissociation. 

Growth is limited by concentration of cells in the medium, which allows easy scale-up.

Can be maintained in culture vessels that are not tissue-culture treated, but requires agitation (i.e., shaking or stirring) for adequate gas exchange. 

Used for bulk protein production, batch harvesting, and many research applications.

Media:

The culture medium is the most important component of the culture environment, because it provides the necessary nutrients, growth factors, and hormones for cell growth, as well as regulating the pH and the osmotic pressure of the culture. Although initial cell culture experiments were performed using natural media obtained from tissue extracts and body fluids, the need for standardization, media quality, and increased demand led to the development of defined media. The three basic classes of media are:-

Basal media,

2. Reduced-serum media, and 

3. Serum-free media. 

which differ in their requirement for supplementation with serum.  

Serum is vitally important as a source of growth and adhesion factors, hormones, lipids and minerals for the culture of cells in basal media. In addition, serum also regulates cell membrane permeability and serves as a carrier for lipids, enzymes, micronutrients, and trace elements into the cell. 

However, using serum in media has a number of disadvantages including high cost, problems with standardization, specificity, variability, and unwanted effects such as stimulation or inhibition of growth and/or cellular function on certain cell cultures.  

Basal Media:

The majority of cell lines grow well in basal media, which contain amino acids, vitamins, inorganic salts, and a carbon source such as glucose, but these basal media formulations must be further supplemented with serum.  

Reduced-Serum Media:

Another strategy to reduce the undesired effects of serum in cell culture experiments is to use reduced-serum media. Reduced-serum media are basal media formulations enriched with nutrients and animal-derived factors, which reduce the amount of serum that is needed. 

Serum-Free Media:

Serum-free media (SFM) circumvents issues with using animal sera by replacing the serum with appropriate nutritional and hormonal formulations. Serum-free media formulations exist for many primary cultures and cell lines, including recombinant protein producing lines of Chinese Hamster Ovary (CHO), various hybridoma cell lines, the insect lines Sf9 and Sf21 (Spodoptera frugiperda), and for cell lines that act as hosts for viral production (e.g., 293, VERO, MDCK, MDBK), and others. One of the major advantages of using serum-free media is the ability to make the medium selective for specific cell types by choosing the appropriate combination of growth factors. 

Advantages:

Increased definition.  

More consistent performance.

Easier purification and downstream processing. 

Precise evaluation of cellular functions. 

Increased productivity. 

Better control over physiological response. 

Enhanced detection of cellular mediators.

Disadvantages:

Requirement for cell type-specific media formulations.

Need for higher degree of reagent purity.

Slower growth.

pH:

Most normal mammalian cell lines grow well at pH 7.4, and there is very little variability among different cell strains. However, some transformed cell lines have been shown to grow better at slightly more acidic environments (pH 7.0–7.4), and some normal fibroblast cell lines prefer slightly more basic environments (pH 7.4–7.7). Insect cell lines such as Sf9 and Sf21 grow optimally at pH 6.2.

CO2:

The growth medium controls the pH of the culture and buffers the cells in culture against changes in the pH. Usually, this buffering is achieved by including an organic (e.g., HEPES) or CO2-bicarbonate based buffer. Because the pH of the medium is dependent on the delicate balance of dissolved carbon dioxide (CO2) and bicarbonate (HCO3 – ), changes in the atmospheric CO2 can alter the pH of the medium. Therefore, it is necessary to use exogenous CO2 when using media buffered with a CO2-bicarbonate based buffer, especially if the cells are cultured in open dishes or transformed cell lines are cultured at high concentrations. While most researchers usually use 5–7% CO2 in air, 4–10% CO2 is common for most cell culture experiments. However, each medium has a recommended CO2 tension and bicarbonate concentration to achieve the correct pH and osmolality; refer to the media manufacturer’s instructions for more information. 

Temperature:

The optimal temperature for cell culture largely depends on the body temperature of the host from which the cells were isolated, and to a lesser degree on the anatomical variation in temperature (e.g., temperature of the skin may be lower than the temperature of skeletal muscle). Overheating is a more serious problem than under heating for cell cultures; therefore, often the temperature in the incubator is set slightly lower than the optimal temperature. 

Most human and mammalian cell lines are maintained at 36°C to 37°C for optimal growth.

Insect cells are cultured at 27°C for optimal growth; they grow more slowly at lower temperatures and at temperatures between 27°C and 30°C. Above 30°C, the viability of insect cells decreases, and the cells do not recover even after they are returned to 27°C.

Avian cell lines require 38.5°C for maximum growth. Although these cells can also be maintained at 37°C, they will grow more slowly. 

Cell lines derived from cold-blooded animals (e.g., amphibians, cold-water fish) tolerate a wide temperature range between 15°C and 26°C.  

Preparation of total cell lysates for immunoblotting

noreply@blogger.com (Anonymous) — Thu, 21 Jun 2018 09:30:00 +0000

The two most common methods of preparing samples for immunoblots are either to lyse the entire cell or tissue mass in sample buffer and load the total cell lysate on the gel or mix any protein sample 1:1 with double-strength sample buffer and load.

Preparing total cell lysates:

In this procedure, samples of cells or tissues are disrupted in an electrophoresis sample buffer. After the chromosomal DNA is sheared to lower viscosity, the samples are processed and run as usual for particular gel system. In the example given  here, the samples are prepared for electrophoresis in a standard Tris/glycinE SDS-Polyacrylamide gel.

Measure the volume of sample to be lysed. For tissue culture cells 109 cells = 1 ml = 1 gram. For tissues or organs, weighing the sample will be sufficient. 1 gram= 1 ml.
Most samples should be boiled for 5 mins. Some antigen will come out of solution under these conditions; if no signal is seen, check different temperature.
Soniate the sample to shear the DNA. Approx. 15-20 sec at full force is sufficient to shear the DNA.
Spin the sample at 10,000 g for 10 mins. Recover the supernatant and discard any pellet. If necessary, determine the relative protein concentration. After the final spin, samples may be frozen at -200 c. They are stable at this step for months.

The samples are now ready for electrophoresis.

To visualize this protocol Click here

References: Antibodies- A laboratory manual; Editor: Ed Harlow and David Lane. Chapter-12 , Immunoblotting, Page no: 480-482.

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Tips for Choosing the Best Antibody

noreply@blogger.com (MedSpace) — Tue, 16 May 2017 02:42:00 +0000

Guides for Selecting the Best Primary Antibody

1. Species

One important factor that should be taken into consideration early in your antibody search is the species of animal in which the primary antibody was generated and whether it will be compatible with your application. If you need to use a conjugated secondary antibody to detect the primary antibody (e.g. when performing immunohistochemistry) the primary antibody should be from a species as phylogenetically distinct as possible from the species of the sample. This will prevent cross-reactivity between the secondary antibody and endogenous immunoglobulins in the sample.

2. The type of antibody matters

Is it better to us a monoclonal or a polyclonal antibody? For some proteins, you don’t have a say in the matter you have to go with whatever antibody is available. However, if you do have a choice, choosing the right kind of antibody might increase your success. Polyclonal antibodies, which recognize multiple epitopes, are often better in Western blots, immunohistochemistry, and immunoprecipitations. They can also be more robust and detect less abundant proteins.

Monoclonals are useful in detecting conformation-specific epitopes and are invaluable when using multiple antibodies simultaneously (such as for detecting two proteins simultaneously in FACS). Due to their specificity, monoclonals often give less background. Because they only recognize a single epitope though, monoclonal antibodies can be more limited in application.

Tips for Selecting the Best Secondary Antibody

Antibody-based assays or immunoassays represent a widely used, valuable tool in areas of basic research, bioprocessing, diagnostics and clinical applications. Although successful detection of the target protein relies on multiple parameters, it is well recognized that the use of high-quality antibodies critically affects assay performance. Because very often immunoassays use secondary antibodies to bind the primary antibody to assist in detection, sorting and purification of target antigens, not only are primary antibodies exhibiting high specificity and sensitivity for the intended antigen essential but also high-quality secondary antibodies are paramount in achieving meaningful results. The following roadmap will guide you step-by-step through the selection process of the best secondary antibody for your experiments:

1. Match the host species of the primary antibody

The first step is to determine the host species that was used to generate the primary antibody. Then, select a secondary antibody specific for detection of the primary antibody species. For example, when using a polyclonal antibody produced by rabbit you will select an anti-rabbit secondary antibody that was raised in an alternative host species such as mouse, goat or donkey. Most primary antibodies are produced in commonly used host species such as rabbit, mouse, goat or chicken. Therefore, anti-mouse, anti-rabbit, anti-goat or anti-chicken polyclonal secondary antibodies are often used for detection. Remember, the species used to generate the secondary antibody should be always different from the host species of the primary antibody.

2. Select the correct reporter based on intended use

Once the source host has been selected, identifying the optimal secondary antibody requires knowledge of the detection assay. For commonly used techniques such as Western blot and ELISA, an enzyme conjugated secondary is most likely the best choice. Good examples are Peroxidase or Alkaline phosphatase. In the case of immunoassays such as immunofluorescence microscopy or flow cytometry (also called FACS) it is more typical to use a secondary antibody conjugated to a fluorochrome (i.e. FITC, DyLight™ or Cy™ dye). For immunoprecipitation experiments, a special reagent that does not detect the precipitating antibody is essential for publication quality images. Rockland’s TrueBlot® products are useful for the accurate detection of secondary antibodies used for immunoprecipitation followed by western blot.

3. Consider using a pre-adsorbed secondary antibody

Pre-adsorption (also cross-adsorption) of the secondary antibody is used to eliminate reactivity from immunoglobulins of undesired species, antibody fragments and/or cell and tissue samples, improving the specificity of an antibody. The degree of cross-reactivity is determined by ELISA or Western Blot detection and is typically less than 1% of the desired signal. The secondary antibody is cross-adsorbed against serum antibody protein from another species or is adsorbed against a mixture of serum antibody protein from several species (i.e., Pre-adsorbed). These highly cross-adsorbed antibodies show low levels of cross-reactivity particularly required in multiple labeling experiments.

4. Define the subclass of the primary antibody

Primary polyclonal antibodies are generated in rabbit, goat, donkey, or chicken and are usually gamma chain immunoglobulins (IgG isotype). Therefore, the secondary antibody should be an anti-IgG antibody that recognizes both heavy and light chain of the primary antibody (anti-IgG H&L). Primary monoclonal antibodies are normally raised in mouse, rat and Armenian hamster but even rabbit and human derived are also used. Because monoclonal IgG antibodies are subclass specific, it is very important you use the secondary antibody directed against that specific subclass. Despite the notion that any anti-mouse IgG should recognize any of the IgG subclasses, recent studies have shown potential bias toward specific subclasses, making the use of anti-mouse IgG subclass-specific essential for “robust and reliable multiplex labeling of target proteins in a variety of applications”. See Manning et al. (2012) for an in-depth review of the subject. When the sub-class of your primary antibody is unknown, you can use anti-IgG F(ab) or consider performing an isotyping assay.

5. Sometimes smaller is better

F(ab')2 fragment secondary antibodies are generated by pepsin digestion of whole IgG antibodies to remove most of the Fc region while leaving the hinge region intact. The resulting fragment is divalent with MW ~110 kDa. Fab fragment secondary antibodies are generated by papain digestion of whole IgG antibodies to remove the Fc region entirely generating a monovalent antibody of ~50 kDa. Both F(ab')2 and Fab fragment antibodies eliminate non-specific binding to the Fc receptors on cells and penetrate tissues more efficiently due to their smaller size. When working with tissues or cells that have Fc receptors (spleen, peripheral blood, hematopoietic cells, leukocytes, NK cells etc.), choose a F(ab')2 and Fab to eliminate non-specific binding to Fc receptors. Fragment conjugated secondary antibodies are ideal for Flow Cytometry, Immunohistochemistry, and Immunofluorescence.

6. Choose the purity level of the secondary antibody

Affinity purified antibodies are isolated by separating monospecific antibodies from other antiserum proteins and non-specific immunoglobulins by solid phase affinity chromatography. Advantages of using an affinity purified antibody include increased specificity, low background, greater sensitivity and lot-to-lot consistency. Affinity purification reduces variation from one product to another, leading to more reproducible immunoassays. IgG fraction antibodies, on the other hand, are very robust and are prepared by a combination of salt fractionation and chromatographic methods with purity and specificity evaluated by different methods. The main benefit of using an IgG fraction is the presence of extremely high-affinity antibodies that may result in a more potent secondary antibody reagent. This may or may not be the cause for affinity purified antibodies which usually exhibit improved specificity sometimes at the expense of affinity. Low abundance proteins or weakly detected primary antibodies detected using an affinity purified secondary antibody may be better recognized using an IgG fraction secondary antibody. Assays, where high background or non-specific binding from the secondary antibody are apparent, may be optimized using an affinity purified secondary.

Conjugate selection guide for secondary antibodies:

Secondary antibodies can be conjugated to a large number of labels, including enzymes, biotin, and fluorescent dyes/proteins. The label of choice depends upon the experimental application.

Immunoassay experiment	Secondary antibody label
ELISA	Enzymes (usually HRP or AP), biotin
Immunofluorescence	Fluorescent label, biotin
Immunohistochemistry	Enzymes, biotin
Flow cytometry	Fluorescent label, biotin
Immunocytochemistry	Fluorescent label, biotin

Enzymes:

Enzyme labels are visualized with chromogenic reactions whereby a soluble colorless substrate is converted to a water-insoluble colored compound. Commonly used enzymes:

Horseradish peroxidase (HRP) is visualized by chromogenic reactions such as diaminobenzidine (DAB) or chemiluminescence. HRP is a 44kDa glycoprotein enzyme label and is more stable than alkaline phosphatase.

Alkaline phosphatase (AP) is a hydroylase enzyme and its signal is often measured through its colorimetric substrate p-nitrophenyl phosphate (pNPP).

Biotin/streptavidin:

Multiple biotin molecules can be conjugated to a secondary antibody, providing an amplification step that makes this conjugate suitable for detecting proteins expressed at low levels. Visualization is through interaction between biotin and streptavidin, where the streptavidin is bound to labels such as HRP or fluorescent probes. During the assay, it is normal procedure for the biotin labeled secondary antibody to be added first, sequentially followed by the streptavidin conjugated to a label.

Fluorescent labeled secondary antibodies:

Fluorescent conjugates are preferred for multi-color analysis. A single dye is excited at a particular wavelength and emits a photon at another wavelength. Alternatively, a tandem dye consists of a donor and acceptor fluorochrome placed in close proximity, allowing energy transfer between the two. The tandem dye is excited at the excitation wavelength of the donor molecule and emits a photon at the emission wavelength of the acceptor molecule.

Tips for choosing a fluorescent labeled secondary antibody:

Choose the brightest set of fluorochromes for your instrument.
Spectral overlap can be minimized through choosing the right fluorescent labels.
Make sure to combine the brightest label with the protein which has the lowest level of expression, and vice versa.
When using secondary antibodies, check to make sure all the primary antibodies have been raised in different species to ensure cross labeling via the secondaries is not encountered. Remember, we provide secondary antibodies against different subtypes from within the same species (mouse IgG1, IgG2, IgG2a, IgG3 etc), providing further choice. Obviously, the corresponding subtype primary must be used.
Pre-adsorbed secondary antibodies are generally used during multi-color analysis to ensure low cross-species reactivity.
Sample autofluorescence is sometimes a problem and must be taken into consideration when choosing your secondary conjugates. For instance, liver sections autofluorescent in the red channel, thus when staining this tissue type labels which emit in the red channel should be avoided.

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Melatonin & Human Sleep

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Egg shell | Industrial applications of egg shell

EGGSHELL CHARACTERISTICS:

APPLICATIONS AND POTENTIAL MARKETS FOR THE DIFFERENT EGGSHELL COMPONENTS:

Stem cells promote peripheral nerve regeneration

Patterning factors | determine regional identity and differentiation potential

Ptf1a | Direct reprogramming of fibroblasts into neural stem cells

RNA isolation from cells | RNA quantification

RNA isolation from cells

Protocol:

RNA quantification

Instruments:

Protocol:

Western blot | troubleshooting | samples turn yellow and precipitate

Why western Blot samples turn yellow and precipitate?

Western blot | Re-use a primary antibody

Why reuse primary antibody?

When reuse is problematic?

When and how we reuse primary antibody?

Western blot | Bradford assay ⎸Protein quantification

A. Preparation of BSA standard curve

Protocol:

B. Protein quantification

Protocol:

Part-1 (Prepare working solution):

Part-2 (Start quantification):

Western blot | ECL development

Protocol

Western blot | Antibody incubation & Washing

Primary & secondary antibody incubation

A. Dilution of primary Ab:

B. Dilution of Secondary AB:

C. Washing:

Western blot | Protein transfer

Preparation:

Transfer protocol:

Western blot | Ponceau S Staining

Ponceau S Staining Solution:

Table-1: Recipe of 0.1% (w/v) Ponceau S in 5% (v/v) Acetic acid

Table-2: Recipe of 0.2% (w/v) Ponceau S in 3% (v/v) Acetic acid

Table-3: Recipe of 0.1N NaOH in 100 ml water

Western blot | Coomassie Blue Staining

Table-1: Recipe of Staining Solution

Table-2: Recipe of Destaining Solution

Protocol

Western blot | Gel Electrophoresis

Preparation of Gel:

Part-1:

Part-2:

Table-1: Acrylamide concentration as per protein size.

Table-2: Preparation of running gel.

Table-3: Preparation of stacking gel.

B. Sample preparation & loading:

Table-4: Special recommendation for sample preparation.

Western blot | Buffer preparation

1. Lysis buffer-cell/tissue

Table-1: Lysis buffer-cell

Table-2: Lysis buffer-tissue

2. Buffer system

A. 30％ Acrylamide solution

B. 1 M Tris-Cl (Lysis buffer pH 7.4 / Stacking gel pH 6.8)

C. 1.5 M Tris-Cl (running gel pH 8.8)

D. 10% SDS

E. 10X SDS running buffer (pH 8.3)

F. 2× sample loading buffer (pH 6.8)

G. 10× Transfer buffer (pH 8.3), 1L

H. 1× Transfer buffer (pH 8.3), 1L

Bradford Protein Assay for determining protein concentration

Principle:

Materials and Reagents

Equipment

Procedure

Microassay procedure (<50 µg ml-1 protein):

Recipes

Cell Culture Environment

Introduction

Adherent vs Suspension Culture:

A. Adherent Culture:

B. Suspension Culture:

Media:

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Relationship between serum irisin and FNDC5 mRNA expression levels.