Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Protocol and Method Guide

Immunofluorescence Microscopy Overview & Theory

Immunofluorescence Microscopy (IF) is a classical technique to observe the localization of molecules in cell/tissue sections. While most researchers try to look for proteins, it is also possible to look for DNA, RNA, and carbohydrates in sections of tissue. With the addition of this technique to your tool belt along with the immunoprecipitation scientific method and the western blot scientific method, you will now have a variety of different ways of manipulating and analyzing biomolecules from tissues, cells, and cell lysates. The principle is fairly straight forward: incubate your sample with an antibody generated towards your target molecule and then detect the antibody using fluorescence. In an immunofluorescence microscopy experiment, this takes the form of putting your cells on a microscope slide, probing with antibodies, and then using specialized fluorescence microscopes with red/blue/green filters along with specific laser emitters to visualize the antibodies. As you would expect, you could either incubate your samples with a antibody-fluorophore conjugate and visualize it (direct immunofluorescence), or you could first put in an antibody that recognizes your target and then put in a secondary antibody-fluorophore conjugate that recognizes the first antibody (indirect immunofluorescence).

Here’s an image which describes the above theory:
Immunofluorescence Microscopy Detect Proteins in Cells
Direct and Indirect Immunofluorescence
Microscopy for localization of DNA and RNA binding proteins

Advantages and Disadvantages of Direct vs. Indirect Immunofluorescence Microscopy

The most common method of performing an IF experiment is to use the indirect immunofluorescence technique. But both methods have their merits, and depending on your application, you may be limited to one method over another.

Direct Immunofluorescence microscopy:
Fewer steps: You don’t need to use a secondary antibody, so you have fewer wash and other steps.
Fewer Complications: Fewer steps also means less troubleshooting.
More expensive: Primary antibodies that are specific and also conjugated to fluorophores are hard to find and expensive.
Lower overall signal: Each primary antibody only has one fluorophore, so you have lower overall signal.

Indirect Immunofluorescence microscopy:
Higher signal: A single primary antibody may bind to multiple secondary antibodies (which each have a fluorophore)–so you get higher overall signal because of this amplification effect.
More versatile: Multiple types of secondary antibodies may be used to detect a single primary antibody. This allows you to probe the same sample multiple times, get higher signal, use a single secondary to detect multiple primaries, and various other strategies.
Cheaper: Secondary antibodies raised against a certain species of primary are very cheap and widely available.
More steps: With a secondary antibody incubation step, there is a possibility for more complications and troubleshooting.

Dealing with the weaknesses of Fluorophores in Immunofluorescence Microscopy

[Slightly technical] Fluorescence is a phenomenon whereby an electron receives some light energy, gets temporarily promoted to a higher energy orbital, and then relaxes back down to its baseline energy state. As the electron relaxes, it releases the light at a slightly lower energy that the initial incident light which hit it. [End Technical Section]

How Fluorescence works:

Immunofluorescence Microscopy Theory of Fluorescence and Electron Energy

This leads to some challenges when performing IF experiments: photobleaching, autofluorescence, and non-specific fluorescence.

Photobleaching during Immunofluorescence Microscopy and how to deal with it:
Use less energy: Some fluorophores will decompose after receiving a lot of energy from lasers. This is typically seen in microscopy when your sample slowly become dimmer and dimmer after imaging a section too long. To overcome this challenge you should use a lower energy intensity when looking around for the right locations in your sample, and then switch to a higher energy intensity when taking an image of your sample. Typically, good microscopy systems will take care of this automatically.

Antifade agents: these molecules scavenge singlet oxygen radicals that are caused by high energy lasers and can be used to maintain high fluorescence signals during microscopy. It is theorized that singlet oxygen species are the main culprit that cause localized damage to fluorophores.

High yield fluorophores: With fewer high-yield fluorophores you can get more signal. The “quantum yield” is an important number when considering which fluorophore you should use alongside your antibodies. If you can find quantum-dot conjugated antibodies, you’re going to be amazed with your microscope images! 🙂 I’ll write more about them in the future.

Cellular Autofluorescence:
There are a few molecules within cells that cause auto-fluorescence. Auto-fluorescence can lead to false positive data in flow cytometry experiments and rarely even in immunofluorescence microscopy. Be wary of the following molecules:

  • FAD and/or FMN: Flavinoids such as these have an Excitation @ 450 nm and Emission @ 515 nm
  • NADH: An Adenine dinucleotide with Excitation @ 340 nm and Emission @ 560 nm

FAD and NADH structures lead to auto-fluorescence:
Immunofluorescence Photobleaching and Autofluorescence

To avoid autofluorescence, use fluorophores with excitations and emissions far away from the above wavelengths so that the inherent fluorescence in your sample doesn’t affect your measured signal.

Non-specific Fluorescence:
Based on the excitation and emission of different fluorophores it is possible for you to get fluorescence signals even though you haven’t probed for them. Be wary of the different fluorophores that you use in your sample and the bandwidths with which you detect them. You may need to modify your microscope’s settings and filters to get really sharp and beautiful IF images.

To avoid non-specific fluorescence, consult a fluorophore chart as mentioned in the guide for the Flow Cytometry (FACS) scientific method.

Why do we have non-specific fluorescence? Take a look at this image:

Immunofluorescence Microscopy Non-specific Fluorescence
Overlapping fluorescence spectra cause problems with Microscopy
Use appropriate and compatible fluorophores

Immunofluorescence Microscopy Step-By-Step Guide

Example Immunofluorescence of Pancreatic Sections
This protocol describes the detection of insulin in paraffin wax-embedded sections of pancreatic tissue.
Normal donkey serum (#D9663 Sigma)
Guinea pig anti-insulin antibody (used at 1:3200; #ab7842 Abcam)
Biotinylated donkey anti-guinea pig antibody (use at 10 µg/ml; ##706-066-148)
Streptavidin conjugated Cy3 (use at 1:100; #016-160-084 Jackson Laboratories)
Proteinase K buffer (50 mM Tris pH 8.3, 3 mM CaCl2, 50% glycerol)
Proteinase K (20 mg/ml; #AM2548 ThermoFisher Scientific)
DAPI (#D1306 ThermoFisher Scientific)
SlowFade mounting media (#S36937 ThermoFisher Scientific)
Experimental procedure:

  1. Cut 8 µm paraffin sections at the microtome and mount onto glass slides.
  2. Heat slides at 60°C for at least 5 min to melt wax.
  3. Dewax slides in xylene for 2x 5min*, 100% ethanol for 2x 5min and immerse in running deionised water until clear.
  4. Circle sections using a wax stick to easily observe staining area in subsequent steps.
  5. Prepare Proteinase K by adding 1 µl to 1 ml warm (37°C) Proteinase K buffer. Perform antigen retrieval by adding 50 µl of diluted Proteinase K per section in a humidity chamber for 20 min at 37°C*. Wash slides 3x 5 min in PBS with stirring.
  6. Block sections with 50 µl of 10% normal donkey serum in PBS at room temperature for 30 min in humidity chamber.
  7. Tip off blocking solution and add 50 µl anti-insulin primary antibody diluted in 10% normal donkey serum in PBS. Incubate overnight at room temperature in humidity chamber.
  8. Wash slides 3x 5 min in PBS with stirring. Add 50 µl anti-guinea pig biotinylated secondary antibody diluted in 10% normal donkey serum in PBS. Incubate 2 h at room temperature in humidity chamber.
  9. Wash slides 3x 5 min in PBS with stirring. Add 50 µl streptavidin-Cy3 diluted in 10% normal donkey serum in PBS. Incubate 1 h at room temperature in humidity chamber in the dark.*
  10. Wash slides 3x 5 min in PBS with stirring. Add 50 µl DAPI (3 µM) diluted in 10% normal donkey serum in PBS to stain cell nuclei. Incubate 30 min at room temperature in humidity chamber.
  11. Tip off excess solution from slide and mount in one drop of mounting medium with coverslip.
  12. Examine at the fluorescence microscope*.

Procedural notes

  • Step 3. Perform in fume hood
  • Step 5. Perform incubations in humidity chamber to reduce evaporation of antibody/proteinase solutions
  • Step 9. Perform incubations with fluorescently labelled reagents in the dark
  • Step 12. Slides can be stored at 4°C until use
  • Consider buying “pre-adsorbed” secondary antibodies. These antibodies have been incubated with common cross-reacting species and they didn’t bind. Therefore, they are super specific to your species of choice and should provide very little background signal.

SciGine Immunoprecipitation Microscopy Protocols and Methods

Immunofluorescence of Arp3
Immunofluorescence Microscopy for MUC of Methanol Fixed Tissue
Use of Cy3 conjugated antibody for Immunofluorescence in Cos7 cells
Cells permeabilized with TritonX-100 for use in Immunofluorescence Microscopy
Analysis of Epithelial Polarization using Indirect-IF


Discussion of Immunofluorescence by Robinson et al.
Robertson et al. IF guide on Biomed Central
NADH and FAD autofluorescence

Immunofluorescence Theory Video by the HHMI

Immunoprecipitation (IP) Scientific Method Guide

Immunoprecipitation Method Guide on SciGine

Immunoprecipitation Overview

Immunoprecipitation is a method for extracting protein from a solution. Typically, this solution is a cell lysate which you want to analyze. Very frequently you’ll hear your colleagues say, «I’m going to do an IP-pull down on my protein» — when you hear this, you’ll know they’re talking about immunoprecipitation. This technique is a must-have for any biochemist who works with proteins because it’s so versatile. Once you remove a protein from solution, you can analyze it to see what it binds. You can also check out its molecular weight and structure. And, beyond proteins, IP can be applied to RNA and DNA pull downs as well.

In theory, this method is very simple. First, lyse your cells using some sort of lysis buffer. Next, add in an antibody that will bind your protein of interest and form a protein-antibody complex. Then drop in some resin which can bind the antibody. Typically, this resin is either agarose-based or superparamagnetic and is covered with Protein A and/or Protein G. These proteins are specifically designed to bind the heavy chains of antibodies so they can easily pull out your protein-antibody complexes. Agarose beads offer higher capacity per bead but magnetic beads are MUCH easier to separate because you can use a magnet to keep them in place. Finally, spin everything down and remove the supernatant. With the remaining bead-antibody-protein conjugates, you can either denature everything and run a SDS-PAGE western blot, or you can try to analyze your protein’s function with an activity assay, or run it on HPLC or LC-MS, etc. There are so many downstream applications of immunoprecipitation!

Take a look at the attached drawing to understand this method:
Immunoprecipitation Scientific Method
Immunoprecipitation Pulldown Assay
Immunoprecipitation IP Downstream applications

Different Types of Immunoprecipitation (IP) / Pull Down Assays

While the previous description shows the most straight forward and common version of immunoprecipitation, there are many variants of this method. It’s truly versatile and powerful, so let’s take a look:

  • Pre-immobilized Antibody Immunoprecipitation: Instead of adding in antibody for your target and then immobilizing the antibody onto agarose beads, you can pre-immobilize it and then add it into your protein mixture.
  • Co-IP, CoIP, or Co-Immunoprecipitation: In a Co-IP, you pull down multiple proteins along with your protein of interest as a complex. It’s very likely that this protein complex that is physiologically relevant to the signaling cascade that they trigger. As an example, you could pull out clathrin-binding proteins and you’d have a clue as to how these proteins enter/exit cells. Cool right? With this simple technique, you can now map out which proteins interact with each other to cause a certain phenotype!
  • ChIP or Chromatin Immunoprecipitation: In this method, the DNA that is bound to your target protein is what you are after. The idea is to first let your protein bind DNA. Then you crosslink it using formaldehyde and lyse the cells. Next, you fragment the DNA so your protein isn’t bound to ALL the cellular DNA…it’s only bound to the DNA that it interacts with. Then do a regular IP based on this blog post. With this precipitated protein, you can then detach the protein from the DNA using heat and perform PCR to look at what DNA segment is being bound.
  • RNA Immunoprecipitation or RIP: Jut like a ChIP, you can also pull out proteins that bind to RNA inside cells. The strategy is the same as ChIP but you need to use RT-PCR to analyze the RNA that you get. We’ll discuss this technique in the future.
  • Biotin/Streptavidin Immunoprecipitation: Some times, Protein A and/or G, based pull-downs don’t work because your antibody doesn’t bind to them very well (due to protein-to-protein variability). In these cases, you need an alternate strategy. If you can find biotinylated antibodies for your target protein, then you can use streptavidin beads to pull them out! Biotin-Streptavidin interactions are among the strongest molecular interactions known to science.
  • Covalent Capture Immunoprecipitation: If you cannot use Protein A and/or G, and you cannot find the biotinylated antibodies for your target protein, then you have to go back to chemistry. The strategy is relatively simple, but most biologists hate chemistry 🙂 All you need to do is to oxidize your antibody to introduce aldehydes in the backbone. This can be done using periodate oxidation on vicinal diols in the heavy-chains of the antibody. Then using amine-containing resins, you can capture the antibodies via schiff base formation and borohydride reduction. You can also flip this strategy on it’s head and use amine-containing antibodies with aldehyde containing resin.

Some of these strategies are shown here:
Preimmobilized antibody or Coip used in IP Pulldowns
Co immunoprecipitation and Chromatin immunoprecipitation
Steps of Chromatin Immunoprecipitation
Finding DNA binding regions of proteins using ChIP

Immunoprecipitation Scientific Method Step-By-Step

Immunoprecipitation of the Lck protein from T cell lysates has been used to analyse the phosphorylation patterns of this important T cell activation protein.

IP Materials:

Lysis buffer (20 mM Tris pH 7.5, 150 mM NaCl, 1% Nonidet P-40, 1 mM phenylmethane sulfonyl fluoride (PMSF))
Protein G-Sepharose beads (Protein G Sepharose 4 Fast Flow #17-0618-01, GE Healthcare Life Sciences)
Protease inhibitors (HaltTM Protease Inhibitor Cocktail 100X, #78430, ThermoFisher Scientific)
Purified mouse anti-human p56-Lck antibody (#551048, BD Biosciences)

IP Experimental procedure:

Perform all steps on ice to avoid protein degradation.

  1. Pellet cells (107) and resuspend in 800 µl ice-cold lysis buffer containing protease inhibitors. Lyse cells at 4°C for 15 min with mixing.*
  2. Pellet cell debris for 15 min at maximum speed on microcentrifuge at 4°C.
  3. Prepare Protein G-Sepharose beads following manufacturers’ instructions, by washing 1 ml of bead slurry three times in lysis buffer. Resuspend the final pellet in an equal volume of lysis buffer and store at 4°C until use.*
  4. Preclear lysate by adding 50 µl of washed Protein G-Sepharose beads at 4°C with mixing for 1 h. Pellet beads for 20 sec at maximum speed on microcentrifuge and retain lysate in a fresh tube.
  5. Add 1-2 µg of specific antibody (eg. anti-Lck) to lysate for 15 min at room temperature. Add 50 µl of washed Protein G-Sepharose beads and incubate overnight at 4°C with mixing.
  6. Spin for 20 sec at maximum speed on microcentrifuge to pellet immunoprecipitates. Wash complexes three times in 500 µl lysis buffer.*

IP Procedural notes:

  • Step 1. Prepare fresh lysis buffer on day of procedure. If protein is intended to be assayed for phosphorylation levels, also include phosphatase inhibitors.
  • Step 3. Protein G has a high affinity for mouse Ig; for other precipitating antibodies Protein A can be used.
  • Step 6. If samples are to be analysed by SDS-PAGE, add reducing sample buffer directly to washed immunoprecipitates prior to electrophoresis.
  • Protein A and/or G are not great way to immobilize antibodies if doing an IP with serum. Serum contains lots of other Protein A and/or G types which will compete with your IP resin.

SciGine Immunoprecipitation Applications

CDK2 Immunoprecipitation
Phosphoserine Pull-down from G418 clones
Immunoprecipitation and SDS-PAGE of FLAG protein
SRC Kinase Immunoprecipitation and SDS-PAGE
Immunoprecipitation of Rheumatoid Arthritis related RV202


Immunoprecipitation guide by Kaboord et. al
IP Protocol by Carey et. al
ChIP by the Haber Lab

Southern Blot Scientific Method Theory and Guide

Southern blot scientific method title

Southern Blots: Overview & Theory

Southern Blotting is a technique that is used to detect whether you have a specific DNA sequence available in your sample. Most commonly, you will be testing for DNA that you get from cell lysates or after creating your own plasmids. You can also do some amazing things like making transgenic mice and proving that you have selected for certain genes of interest to you! All you have to do is Southern Blot any set of DNA from the mice and you’ll have concrete proof. As you probably understand, this is one of the most common techniques that geneticists and molecular biologists know because everything they do needs to be proved by a southern blot.

To execute a southern blot, first collect some DNA. This can be from a cell lysate or from tissue samples, etc. Next, digest the DNA using DNAse and run these fragments on an agarose gel. This will separate the DNA by size and you’ll know which well in the gel corresponds with which sample. Next, in a similar fashion to Western blots, this DNA is then transferred onto blotting paper (typically made of nitrocellulose or nylon). After the “blotting” process is complete, the DNA is then “probed” using a radioactive (or fluorescent) DNA sequence that is complementary to the sequence that you want to detect. Finally, this “probed” radioactive blot is then imaged using an autoradiograph. The steps for this process are described in the following image:
Southern Blot Scientific Method
Southern Blot Step by Step
Southern Blot Tips and Tricks
Southern Blotting Method

Nuances of Southern Blotting theory

Why is there is a second denaturing step for DNA that’s 15 KB or larger?
It’s because these large pieces of DNA don’t transfer very well to the blotting paper. As you can imagine, the larger a piece of DNA is, the slower it will migrate through an agarose gel. Even after leaving the blotting paper overnight, the transfer may not be complete!

How does the DNA actually go from the gel into the blot?
Through capillary action and wicking! Unlike a western blot where a voltage gradient is utilized to pull proteins into the blotting paper, in Southern Blots, the DNA merely moves over to the blotting paper overnight without much force at all. That’s why it is incredibly important to make sure that the blotting paper and the gel are in close contact. It’s also very important to make sure that the glass plate on top is heavy enough so that it forces the gel and the blot together.

Why is there a UV step?
By using UV after the transfer step, the DNA (which has some free aldehyde groups due to depurination) can react with the nitrocellulose/nylon membrane to form covalent bonds.

What do the NaOH and the HCl do during the denaturation step?
Essentially HCl removes some/all of the purine bases from the DNA and makes the two DNA strands less sticky to each other (because there is less hydrogen bonding). This process is called depurination. NaOH also prevents the two strands from forming hydrogen bonds due to deprotonation of all bases.

How does radiolabeling with P32 work?
Small amounts of DNAse introduce nicks into the single stranded probe DNA. DNA polymerase then utilizes the dATP32 from solution to repair these nicks and incorporates these radioactive phosphates into the backbone of the DNA.

Southern blot Step-By-Step Guide

Materials for Southern Blot of Mouse Tail DNA

Denhardt’s Solution 50X (5g Ficoll 70000, 5g Polyvinyl pyrrolidone, 5g BSA Fraction V in 500 ml water)
Hybridization cocktail (25 ml 50% dextran sulfate, 25 ml 20X SSC, 50 ml formamide, 1 ml Tris 1M, 2 ml Denhardt’s Solution 50X, 1ml 10% SDS) – prefilter before use
TE buffer (10 mM Tris HCl pH 7.5, 1 mM EDTA)
BamHI enzyme and buffer (#R0136S New England BioLabs)
100X BSA (B9000S New England BioLabs)
Gel loading dye (#G2526 Sigma)
TBE buffer 10X (#93290 Sigma)
SSC buffer 20X (#S6639 Sigma)
[α32P]dCTP (3000 Ci/mmol; #NHG013H250UC Perkin Elmer)
Sephadex G-50 (#G50150 Sigma)
Ready-To-Go DNA Labeling Beads (-dCTP) (#27-9240-01 GE Lifesciences)
Whatman paper (#WHA10427810 Sigma)
Salmon Sperm DNA (#15632011 ThermoFisher)

Southern Blot Protocol
DNA digest and gel electrophoresis

    1. Setup DNA digest reactions from collected tail samples as follows:

10 µl DNA (12 ug)
6 µl 10X BamHI buffer
0.6 µl 100X BSA
1 µl BamHI
42.4 µl H2O
Incubate at 37°C for at least 5 hours.

  1. Add 6 µl gel loading dye to each sample.
  2. Run samples of a 0.8% agarose gel in TBE for 16 h at 25V.
  3. Stain gel in 1 µg/ml ethidium bromide for 30 min.
  4. Take a photo of the gel.*


    1. Soak gel in 0.2 N HCl for 10 min with shaking to depurinate (remove purines from DNA). Rinse with water.
    2. Soak gel in two washes of 500 ml 0.4 M NaOH/1.5 M NaCl solution for 30 min each time.
    3. Soak gel in two washes of 500 ml 1 M Tris/1.5 M NaCl solution for 30 min each time.
    4. Setup transfer stack by placing 2 layers of Whatman paper on glass plate (wet before laying down) with soaked gel on top (flipped over). Cover with plastic wrap.
    5. Using a razor blade, cut out plastic over gel and place wetted membrane on gel. Place two layers of Whatman paper on top (wet before laying down).
    6. Place a stack of paper towels on the top and cover with a glass plate and transfer in 6X SSC overnight.
    7. Dismantle and air-dry membrane.
    8. Expose membrane to UV for 90 s (DNA side down). Bake between Whatman paper at 80°C for at least 1 h.

Labelling the probe

      1. Denature probe by adding 2 µl probe (25-50 ng) to 44 µl TE and incubating at 95°C for 4 min). Immediately place on ice.
      2. Add 46 µl denatured probe and 4 µl of [α32P]dCTP to Ready-to-Go labelling bead tube. Flick to mix.
      3. Incubate at 37°C for 15-30 min.
      4. Add 50 µl TE and run through a G50 sephadex column

Probe Hybridization and Autoradiograph

      1. Add 50 ml hybridization cocktail to the membrane.
      2. Add 300 µl of 10 mg/ml salmon sperm DNA to labelled probe and denature at 95°C for 5 min. Immediately place on ice. Add to membrane and cocktail solution.
      3. Hybridize overnight at 42°C.
      4. Wash briefly with 50 ml wash solution (0.2X SSC/0.1% SDS).
      5. Wash for 30 min with 100 ml wash solution at 50°C. Repeat.
      6. Check blot with Geiger counter to estimate cpm.
      7. Expose membrane to film and store at -70°C.
      8. Develop after 2-5 days.

Notes on this Southern Blot method

      • During DNA digest Step 5 you can note the position of the marker bands by using a fluorescent ruler or marking bands directly on the gel by punching small holes with a needle.
      • This method uses dCTP32 and not dATP32 as we talked about in the theory section
      • After undergoing HCl and NaOH treatment for the denaturation step, it is very important to neutralize the gel once again.
      • If you don’t want to buy the labelling beads, an alternative strategy is presented here at MIT Cores

Application of Southern Blots on SciGine, a Scientific Method Search Engine


ASU Southern Blots Guide
MIT Core Guide
DNA Blot from Davidson College

Southern Blot Video by Shomu’s Biology

Western Blot Theory and Method Guide

Western Blot Method Guide and Step by Step Procedure

Overview of Western Blot Method

A western blot enables sensitive detection of specific proteins from a solution containing multiple proteins. This is an essential biology technique and one of the cheapest methods that can be utilized to analyze proteins. To perform a western blot first separate proteins based on their mass and charge via gel electrophoresis, and then follow up by detecting the protein of choice with a specific antibody. Typically, researchers will use western blots to separate proteins from cell media or from cell lysates. For example, if you wanted to find out how much actin your cells are expressing, a western blot can easily compare actin amounts between different cell types. It’s also likely that you will be using western blots when producing proteins in mammalian and insect cells.

In a typical western blot procedure, cells will first be lysed and the amount of protein will be determined using a spectrophotometer. Then a gel will be made and the total protein from the cell lysate will be loaded into wells in the gel. After applying an electrical field, the proteins in the gel will begin to migrate down and separate into distinct bands based on the size and charge of the protein. After the smallest proteins reach the bottom of the gel, the electrophoresis will be stopped and all proteins on the gel will be transferred onto blotting paper so that they can easily be handled. Finally, antibodies that recognize the proteins of interest will be added and detected via chemiluminescence.

Here is a step by step illustration of how to perform a western blot:

Western Blot Scientific Method Guide

Western Blot Step By Step

SDS-PAGE Western Blot Step-by-Step Protocol

Western blotting can be used to examine the upregulation of RCAN1, a signaling molecule in neuronal cell types.

Materials for Western Blots:

  • Rabbit anti-RCAN1 antibody (#SAB2101967, Sigma-Aldrich)
  • SDS-PAGE gel (Criterion TGX precast Stain-free Any kD gel, #5678124, Bio-Rad)
  • TBS (20 mM TrisCl pH 7.6, 150 mM NaCl)
  • Running buffer (25 mM Tris, 192 mM glycine, 0.1% SDS)
  • Transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol, 0.05% SDS)
  • 4X SDS-PAGE loading buffer (Laemmli’s sample buffer #1610747 Bio-Rad; add fresh dithiothreitol to 10 mg/ml on the day of experiment)
  • Transfer membrane (0.2 um polyvinylidene fluoride membrane, #03010040001 Roche)
  • Secondary antibody (donkey anti-rabbit horseradish peroxidase-conjugate; #711-035-152 Jackson ImmunoResearch)
  • Blocking buffer (5% skim milk powder in TBS with 0.1% Tween20)
  • Immun-Star WesternC Chemiluminescence reaction solutions (#170-5070 Bio-Rad)
  • Dual color ladder (#1610374 Bio-Rad)
  • Blotting paper (ProteanXL, #1703966 Bio-Rad)

Western Blot Experimental procedure:

  1. Unwrap precast gel and rinse wells three times with running buffer. Assemble gel in tank and fill with running buffer.*
  2. In an Eppendorf tube add protein sample (30 µg) to 10 µl 4X SDS-PAGE loading buffer and add water to a final volume of 40 µl.
  3. Heat samples to 95°C for 2 min and spin briefly to ensure contents are at the bottom of the tube
  4. Load gel with samples and include ladder in one lane.
  5. Run gel at 200V for 30 min.
  6. While gel is running, soak two pieces of blotting paper (cut to the same size as the gel) in transfer buffer (approx. 30 min). Activate transfer membrane (also cut to size) by dipping in methanol, then soak in transfer buffer for approx. 10 min.
  7. Remove gel from tank and place in transfer buffer.
  8. Assemble transfer “sandwich” by placing down soaked blotting paper, transfer membrane, gel and blotting paper onto open transfer cassette (Turbo Blot transfer unit; Bio-Rad). Use a glass rod to roll across the “sandwich” to remove any air bubbles.
  9. Close cassette and run in machine (standard minigel program for 30 min).
  10. Remove transfer membrane from cassette, taking care to snip one corner to ensure orientation.*
  11. Incubate transfer membrane in blocking buffer for 1 h at 4°C with rocking.
  12. Pour off blocking buffer and add diluted anti-RCAN1 antibody 1:200 in 5 ml TBS with 2.5% skim milk power and 0.05% Tween20. Incubate overnight at 4C with rocking.
  13. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time.
  14. Incubate with diluted secondary antibody 1:2500 in 5 ml TBS with 0.2% Tween20 at 4°C with rocking for 1 h.
  15. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time. Rinse membrane briefly in water.
  16. Mix 1 ml each of ECL reagents in a foil-wrapped tube and add to membrane for 5 min prior to imaging on ChemiDoc MP imager (Bio-Rad).

Procedural notes for this Western Blot Method:

  • This precast gel contains 18 lanes with a loading capacity of 10-40 ug protein in up to 30 ul per well
  • Small needle-point markings can be added to membrane in-line with color markers which reduce in intensity following subsequent incubation and washing steps.
  • To make sure you know which step you are on, cut the bottom right side of your gel after running the gel electrophoresis.
  • In this method, the protein is denatured prior to running on the gel. This is called SDS-PAGE. By denaturing, you ensure that the size and charge are all that matter, as opposed to native gel electrophoresis where the conformation of the protein also matters.
  • Blots can be regenerated (the antibodies that were used for probing can be removed) by using stripping buffer. However, blots can only be stripped a few times before they have too much background noise to be easily analyzed.

Applications of Western Blots on Scigine (Search Engine for Scientific Methods):


NIH Western Blot Reference
Western Blotting by Kurien et al

Good Video References Related to Western Blots:

Flow Cytometry and FACS: Method Guide

FACS & Flow Cytometry for Cell Population Analysis

FACS Method Overview: Using Fluorescence to Understand Cells and Cell Populations.

FACS, or Fluorescence Assisted Cell Sorting, is a type of technique that enables you to understand cells by tagging them with fluorescent markers. There are a number of things that FACS allows you to do:

  • Separate one type of cell from a mixture of cells
  • Count how many cells are in a mixture
  • Detect what kind of biomarkers a cell might have (is it cancerous? Just a regular lymphocyte? Etc.)
  • Find out if your cell is making a protein of choice (similar to biomarker detection)
  • And more.

It’s an amazing and powerful technique that you should always keep as part of your skills. Most Bio-Pharmaceutical companies have FACS experts in-house because it is so versatile!

The principle of FACS is simple:

  1. Label cells of interest with a marker such as an antibody
  2. Pass the cells through a laser and detect which ones have the antibody
  3. Separate those cells from the rest

Here’s what this looks like in a simple picture:
FACS and Flow Cytometry Method Overall

What does the FACS instrument detect?

  • Forward Scatter Light (FSC): When you shine a laser on an object, the object blocks some of it and the rest continues to go forward towards the detector. This is called forward scattered light.
  • Side Scatter Light (SSC): An object might also bounce some of the laser in an alternate direction. There is a detector that is orthogonal to the light direction and detects this scattered light.
  • Fluorescence Emission Signal: Excite your fluorophores with a certain wavelength of light and they’ll emit a different wavelength depending on the type of fluorophore. Fluorescence detectors in the instrument will detect this emitted light.

Take a look at this illustration:
FACS Detection of Molecules and Cells
Based on the previous section, FACS provides data on forward scattered light, side scattered light, and fluorescence intensity. By graphing these different values together, you can get an idea of what your cell sample is like. Here’s the kind of data you would get:

  • Plot of FSC vs. SSC:
    • Cell debris in your solution is usually small, so it would have low FSC and SSC values.
    • Normal cells have medium sized FSC and SSC values
    • Large cells like Granulocytes have large FSC and SSC values
  • Plot of Fluorescence intensity:
    • After cleaning your cell solution and removing any non-specific fluroescence, your cells will be the only thing with large fluorescence intensity
    • A plot of intensity provides you a histogram telling you how much fluorescence is in the solution
    • You can use this to find out how many cells are there, how many antibodies bound to each cell, how much of a certain biomarker is there, etc.
    • You can compare different fluorescence histograms to compare cell types as well

Take a look at these images:
FACS Understanding Dot Plots and Histograms

Gating: How to measure cells of only one population and ignore debris

In the above images you can see that your figures may provide information on several types of particles: debris, small cells, large cells, particles with high and low fluorescence, etc. In order to narrow down your results, most instruments allow you to “Gate” your results so that the instrument provides you with a new graph within limits that you set. As an example, you may plot FSC vs. SSC and then gate only for larger particles. Then using that gate you may look at the fluorescence intensity in Red, Green, and Blue. This way you can analyze cells that have red biomarkers, or cells that have blue biomarkers specifically.

As an example, what if you wanted to only collect cells that highly express one cancer-related cellular protein (ex: folate receptor) ?

Using FACS, this would be easy to analyze! Just mark your cells with an anti-folate receptor antibody with GFP. Then, use FACS to separate out the cells that express the receptor in your sample. Take a look at this image to understand this concept.
Gating in Flow Cytometry

Complete Step by Step Method Guide

Unfortunately, this is such a highly instrument-oriented technique that it’s difficult for me to provide details on how to operate a FACS machine. Nonetheless, take a look here to get an idea.

Tissue Culture Flask
With cells such as HT-29
Trypsin such as Tryp LE from Invitrogen
Serological pipette
Without pH indicator so that there isn’t any extraneous fluorescence


  1. Grow HT-29 to confluence in a T-75 flask in 7 ml of DMEM
  2. Use lab protocols to mark them with any antibodies that you’d like such as an anti-folate receptor antibody
  3. Aspirate the DMEM and add PBS 1x
  4. Swirl around and remove PBS
  5. Wash once again with PBS by following the previous 2 steps
  6. Add 2 ml of Trypsin (TrypLE) and wait until cells detach for approx 10 minutes
  7. Collect cells into a falcon tube by using a serological pipette
  8. Use a hemacytometer to dilute cells until they are at a concentration of 1×106 cells/ml using DMEM
  9. Load cells into the FACS using using appropriate 5 ml tubes
  10. Make sure the instrument is running well by using control samples or any extra cells that you have
  11. Replace cell collection tubes and make sure they are sterile if you plan on culturing cells that are collected
  12. Run the FACS instrument 🙂

Notes on this FACS/Flow Cytometry Methodology

  • FACS may also be referred to as Flow Cytometry on Job Postings. This is because FACS is a part of the overall group of techniques called Flow Cytometry. In biology, however, it is unlikely that you will use any other techniques besides this one.
  • The types of tubes that are necessary for loading a FACS unit vary between instruments
  • Some cells clump up and make data analysis harder. Try diluting them more or mixing them more using your serological pipette
  • It’s important to set the instrument’s gain settings to maximize/minimize signals as appropriate
  • Make sure to also set the correct excitation/emission levels for your various fluorescence markers. You will also need to set your thresholds such that your signals don’t overlap.
  • If you plan on using multiple fluorescence markers at the same time make sure you consult a fluorescence ex/em table like this fluorescence spectra table. You need to make sure that your fluorescent signals don’t overlap.

Applications of FACS on SciGine

Analysis of cell cycle and viability with a FACScan
Analysis of a GFP reporter gene using FACS
Analysis of gene disruption in diploids
Using FACS to analyze camptothecin induced apoptosis
FACS analysis of cell cycle using Propidium Iodide


Excellent Antibodies-Online FACS reference
Thermofisher FACS manual
What is FACS from UMass

HPLC: Biochemical Analysis. A Step-By-Step Method Guide

HPLC Analysis Step by Step

HPLC Method Overview

HPLC, or high performance liquid chromatography is an amazing analytical technique for chemical compounds including biopolymers, small molecules, and polymers. In this method, a sample is first dissolved to make a solution. This solution is then injected into a “column” that contains resin that will interact with the sample. This will slow down the movement of the sample through the “column” and as the sample comes out the other side of the column, it is detected. This allows you to know both the time at which the sample comes out and the intensity of the sample that was detected. Here’s an overview of this technique:

HPLC Bioanalytical Method Guide

So, while there is continuous flow of some buffer through the column, we also inject our sample and observe as different molecules within the sample come out at different “retention times”. The detector on the end of the column can be any kind of detector but the most common types are refractive index (RI), ultraviolet (DAD), and fluorescence (FLD). Each of these will detect different properties of the molecules that come out of the column and display a chromatogram.

HPLC Chromatogram Guide

Types of Chromatography

Different column resin compositions determine the kind of chromatography that you are running and what molecules you can separate.

  • Normal Phase: The column is filled with silica particles which are polar and the buffer running through the system is non-polar. Once you inject your sample, polar particles will stick to the silica more and have a longer retention time than non-polar molecules.
  • Reverse Phase: The column is filled with hydrophobic particles (actually they are silica particles with long hydrocarbons on the surface). The buffer that is running through the system is polar (such as acetonitrile/water or methanol/water mixtures). This means that hydrophobic molecules will stick to the resin more and be retained longer.

Complete Step by Step HPLC guide


HPLC autosampler vials I only use autosamplers since manual injection is tedious 🙂
Centrifugal filters with 0.2 um pores To clean up samples
Eppendorf vials For centrifuging
HPLC machine


In a typical HPLC procedure you can decide the following variables:

Variable What it does
Flow rate With fast flow peaks come out sooner but there’s they’re harder to resolve and tend to blend together. For more resolution, run slower.
Pressure Affected by flow rate and solvent
Solvent Buffers Determines signal intensity, how quickly the peaks come out, signal fidelity
Column Type Determines the type of interaction with the sample
Detection Parameters If using UV or FLD, you need to set the right excitation/emission wavelengths

Since HPLC is a very machine-variable technique, I can only provide general guidelines.

For sample preparation:

  1. Dissolve your biopolymers or small molecules in a suitable solvent such as methanol
  2. Centrifuge at 10,000 rcf in an eppendorf vial and keep the supernatant to remove any large particular matter
  3. With a centrifugal filter, add 500 ul of your sample solution onto the top
  4. Centrifuge at 10,000 rcf and collect the filtrate (the solution that successfully passes through the filter)
  5. Load this sample into an HPLC vial

For setting up the HPLC machine:

  1. Make sure you have all your buffers set up
  2. Open the purge valve and purge the system for 5 minutes.
  3. Add your samples into the autosampler tray
  4. Stop the purge
  5. Close the purge valve
  6. Run the system at a normal flow rate (1 ml/min) with your buffer to equilibrate the column for 10 minutes
  7. Make sure that your pressure is stable (ie, less than 2-3 bar of fluctuation)
  8. Set up your sequence and your method
  9. Run a standard before your actual samples or as part of the same sequence

Example buffer system to determine Fluoresceinamine levels in samples:
Sample: Add 10 ug of fluoresceinamine into 1 ml of Acetonitrile.
Buffer: Pure acetonitrile buffer on a C-18 column; this is “reverse phase”.
Flow rate: 1 ml/min.
Column: 4.6 mm x 30 cm size.
Detection: Detect via a fluorescence detector set to Excitation @ 485 nm and Emission @ 535 nm.

Notes on HPLC methodology

  • To clean the system and equilibriate it, you need to run enough solvent. However, this amount varies column-to- column. A typical 4.6 mm x 30 cm column should be clean when you follow the procedure above.
  • Isocratic means that the solvent concentration stays constant throughout the run.
  • It is useful to run standards before your samples as well as with your samples. Standards make it easy to identify which peak pertains to your molecule of interest.
  • Always use HPLC grade solvents. This is especially true for solvents like THF which are frequently sold with inhibitors that also complicate your ability to detect your molecule of interest.

Applications of HPLC on SciGine

HPLC is such a versatile technique. Take a look at these methods on SciGine which assay different types of chemicals in various samples.


ChemGuide Summary of Technique
Method Guide from Waters
Overview on Wiki

Using PCR To Amplify DNA, A Step-By-Step Guide

PCR Method Guide on SciGine

PCR Biological Method Overview

PCR, or polymerase chain reaction, is a method to amplify a segment of DNA for analysis. Because it is such a powerful technique, there are a HUGE number of situations where PCR may be used. Some common reasons for using it are:

  1. Microbiology: You need to know if your bacteria was transformed properly with your plasmid
  2. Oncology: You need to know if a particular gene exists in your cancerous cells
  3. Forensics: You need to know if a certain criminal’s DNA was present at the crime scene

The basic steps of PCR include:

  1. Designing primers to designate a target DNA sequence to amplify
  2. Mixing together the primers with the target DNA strand, polymerase enzyme, and deoxynucleotides
  3. Running a thermocycler multiple times in “cycles” to repeatedly:
    • Separate DNA strands
    • Allow primers to anneal
    • Allow the polymerase to attach and synthesize a new DNA chain
  4. Sequencing or Electrophoresis to prove that the PCR worked

In a nutshell, here is what Step 3, above, looks like:

PCR, a biological method, amplifies DNA - SciGine

Ingredients in a PCR mixture – What every component does

In general, a PCR tube will contain the following items:

Water: Solvent
dNTPs (or deoxynucleotide triphosphates): Single bases A, T, C, and G which are used by the polymerase while replicating the DNA. As the polymerase adds base pairs onto the new DNA strand, one base pair is used at a time.
MgCl2: An essential cofactor for the polymerase enzyme
Primers: Short segments of single-stranded DNA used to frame the DNA region that needs to be amplified. They are complementary to the template DNA strand only at defined locations around the target sequence.
Target DNA: The DNA “template” that you want to make copies of. This can be a full DNA chain or a part of a longer chain.
Taq Polymerase: An enzyme from Thermis aquaticus that uses dNTPs and replicates DNA starting from the 3′ end of a template strand towards the 5′ end. Taq  is used in PCR specifically because it is resistant to the high temperatures used for separating DNA strands during Step 3a above .

Complete Step-By-Step PCR Protocol


PCR tubes

PCR temperature Cycler


Reaction Buffer

10 mM Tris-HCl, 50 mM KCl, and 1.5 mM MgCl2, pH 8.3


  1. For each PCR tube set up the following mixture of materials up to a 50 ul total volume on ice

    dNTP solution

    200 uM for each dNTP

    Forward Primer

    0.2 uM

    Reverse Primer

    0.2 uM

    Target DNA

    Less than 1000 ng

    Taq Polymerase

    1.25 units per tube

    Nuclease Free Water

    Q.s up to 50 ul

    Mineral Oil

    Add a little bit on top of each tube if you don’t have a heated lid on the temperature cycler

  2. Pipette each tube up and down several times, gently, so as not to add bubbles
  3. Centrifuge each tube down for 1-2 seconds at 100 rcf to bring all contents to the bottom
  4. Heat up the temperature cycler to 95oC
  5. Quickly transfer tubes from ice to the temperature cycler and begin thermocycling
  6. Typical thermocycle procedure:

    Step Name




    95 oC

    30 seconds

    Cycle Denaturation

    95 oC

    30 seconds

    Cycle Priming

    50-60 oC

    60 seconds

    Cycle Extension

    72 oC

    1 minute per kilobase of target DNA

    Final Extension

    72 oC

    5 minutes

    Hold Temperature

    4 oC


  7. Repeat the above “cycle” steps 35-45 times.
  8. Make sure to keep the samples on ice while you are not using them. The infinite hold sequence allows you the flexibility of leaving your samples in the temperature cycler while you are busy with other lab work.

Notes on this PCR Methodology

  • Note 1. It is important to design your PCR primers to be specific to only the regions flanking the target sequence. Typically, specific primers are ~30-40 bases in length.
  • Note 2. The Tm (melting temperature) of the primers affect the temperature in Step 3b and the “Cycle priming” step.
  • Note 3. For greater accuracy/fidelity while copying DNA, a Pfu polymerase (from Pyrococcus Furiosus) may also be used.
  • Note 4. With 35 cycles, the target DNA is amplified 236 times its initial concentration. For times when you have very little target DNA, you can amplify 45 times or more.
  • Note 5. For Primer design, you can use tools such as Primer 3. Tools are also available for calculating the Tm of your primers such as this Tm calculator.
  • Note 6. G-C rich sequences will need longer denaturation times (typically up to 5 minutes) because of the additional hydrogen bonding. Any template with more than 60% G-C bases is considered G-C rich.

ELISA: A Step By Step Method Guide

Step by Step ELISA Guide on SciGine

ELISA: A Step By Step Method Guide

ELISA Biological Method Overview

ELISA is the common acronym for Enzyme-Linked-Immunosorbent Assay. It’s a quick plate based technique for detecting an antigen from a solution. This antigen could be a peptide, protein, antibody, or small molecule. In general, for an ELISA, an antigen is first immobilized on a surface (Step 1 below). Next, an antibody specific to the antigen is flowed over the surface (Step 2). This antibody, is also attached to a chemiluminescence-related enzyme. Treatment with the chemiluminescent substrate facilitates detection of the antibody and the antigen (Step 3). Take a look at these pictures to get an overview of the strategy:

ELISA Steps - SciGine Biological Methods

Types of ELISAs

There are a few different types of ELISA assays but they all follow the basic strategy outlined above.  Essentially, one can choose how to immobilize the antigen on the surface and how the antigen is  detected via the antibody.

  1. Direct Assay: In this method, the antigen is immobilized to the surface and detected directly via an  antibody that’s bound to a chemiluminescent enzyme. (Same as above)
  2. Indirect Assay: In this method, the detecting antibody doesn’t have the chemiluminescent  enzyme. So, another antibody must bind to the first antibody to facilitate detection.
  3. Sandwich Assay: The most common type of ELISA. In this assay, a “capture” antibody is first  immobilized to the substrate. Then antigen is flowed over it so that it gets immobilized to the  surface along with the capture antibody. Finally the detection antibody is flowed over the  substrate and it binds the antigen. This detection antibody may be directly conjugated to the  chemiluminescent enzyme (just like a direct assay) or another antibody may be needed (just like  the indirect assay).

Types of ELISA Assays - SciGine

A Complete Sandwich ELISA protocol

Materials for ELISA

96 well polystyrene plate

Plate shaker


Coating buffer

0.2 M sodium carbonate/bicarbonate buffer, pH 9.4

Wash buffer

0.1 M phosphate, 0.15 M sodium chloride, pH 7.2 with 0.05% Tween 20

Blocking buffer

2% w/v Bovine Serum Albumin in Wash Buffer

Diluent buffer

2% w/v BSA in Wash buffer or a more appropriate buffer such as cell culture media

Stop buffer

2 M Sulfuric Acid

Capture Antibody Solution

15 ug/ml antibody in coating buffer

Detection Antibody Solution

10 ug/ml in (20% Diluent buffer/80% Wash Buffer)

Enzyme Conjugated Antibody Solution

200 ng/ml in (20% Diluent buffer/80% Wash Buffer)

HRP Substrate

TMB (3,3′,5,5′-tetramethylbenzidine). 1 mg/ml. Usually commercially available as a solution.

Step-By-Step Method for ELISA

  1. Prepare a standard curve with your antigen in Diluent Buffer spanning a wide range of concentrations from 0 pg/ml to 3 times your maximum expected antigen concentration (3000 pg/ml approximately)
  2. Dilute the capture antibody to 15 ug/ml and have enough for 100 ul/well
  3. Add the capture antibody to the polystyrene plate, cover, and incubate at room temp. for 2 hours
  4. Remove the solution from each well and add in wash buffer (200 ul per well). Shake for 5 minutes. Repeat 3-5 times.
  5. Add 200 ul of blocking buffer per well, cover and incubate at room temperature for 1 hour (or overnight at 4 oC).
  6. Prepare the samples and standards such that you have 100 ul per well
  7. Remove the wash buffer and add in your sample + standard antigens into different wells. Cover and incubate at room temperature for 1 hour
  8. Repeat Step 4  to wash the plate
  9. Add 100 ul of the Detection Antibody per well. Incubate at room temperature for 1 hour.
  10. Repeat Step 4 to wash the plate
  11. Add 100 ul of the Enzyme conjugated Antibody to each well and incubate for 1 hour at r.t.
  12. Repeat Step 4 to wash the plate (2 times). We need to make sure the plate is very clean and any non-specific binding is minimized.
  13. Add 100 ul of the HRP substrate solution (1 mg/ml TMB)
  14. Incubate until blue (usually about 10 minutes at room temperature)
  15. Add 100 ul of Stop Buffer. This should make the solution yellow.
  16. Measure using a plate reader at 450 nm absorbance.

Notes on this ELISA method

      Note 1. Your standard curve needs to span beyond your antigen concentration because you need to determine the exact amount of your antigen within the linear range of the standard curve. If necessary, dilute your antigen solution down to a point where it is within your standard range.
      Note 2. Concentration of antibodies used will need to be optimized. It is highly likely that you will need to dilute each of the antibodies down rather than increase their concentration because these are at the upper ranges of the necessary concentration.

Tools for Scholarly Search & Research in Biology

Efficient scholarly research helps to reduce unnecessary experiments

Scholarly Research reduces unnecessary experimentation

Nobody wants to reinvent the wheel. Look at all of the pain staking work that goes into re-testing previous experiments and developing methods that have already been known for decades. By taking our time to define our problems, hypothesize our solutions,and search through literature prior to executing on our plans, we can dramatically improve the effectiveness of our research. In fact, people dedicate entire chunks of their lives to being good at searching and testing! (Hint…it’s called a PhD).

In this blog post, I wanted to discuss the tools that are available to help Biologists and biochemists perform effective research. In particular I wanted to focus on biology-related search engines and their development/features since their inception.

A list of tools for biologists to efficiently perform their scholarly searches

  1. Scigine: A search engine for methods in biology, biochemistry, pharmaceutics, and clinical science. Provides step-by-step methods along with the ability to take “notes“, modify methods, and view them on a phone/tablet while performing experiments. Users can also share their notes with colleagues with the click of a button. (~600,000+ methods)
  2. A “git-hub” for biology with the ability to “fork” methods and modify them. Also includes the ability to form groups with others to share methods online.  (~700 methods)
  3. Protocols-online: A collection of methods aggregated from multiple authors along with forums to ask questions and trending jobs for biologists and biochemists. (~1000 methods)
  4. Vadlo: Or “Fig”, hones google in to specific biology-related websites by providing a custom search text box. It is possible to use this search feature to broadly find methods, presentations, and articles that would normally be found through google.
  5.  DOAJ: The directory of open access journals provides access to over 1.9 million journal articles in biology, biochemistry, and related fields. In case your library does not provide access to journal articles, the DOAJ is a solid resource that is freely available.
  6. PubMed: From the website – “PubMed comprises more than 26 million citations for biomedical literature from MEDLINE, life science journals, and online books. Citations may include links to full-text content from PubMed Central and publisher web sites.” This is the go-to site for scholarly research in biology and related fields.
  7. Microsoft Research Tools: A collection of online tools for genomics and bioinformatics provided by the Microsoft Biology Initiative.
  8. PubAlert: Automatic weekly or daily digests of PubMed searches for your keywords. This is a great tool for keeping up with the latest research in your field. Example, setting up an alert for “Actin” will provide you with an email digest of the most recent articles with “Actin” in the title or abstract.

Hopefully, this blog post provides a good “lay of the land”.

Enjoy the list of tools!


SciGine: Inception of A Biology Methods Search Engine

Biology and Biochemistry Methods Search EngineDoing a PhD is about the pursuit of knowledge

The typical biology PhD student spends 5+ years in a PhD program. If you were to follow a student, you might start in the morning by watching him/her looking through papers to check what latest research might be out there related to their topic. Have they been scooped? Is there contradictory research out there that they now have to mention in their next research paper? What new information can they glean from their PubMed searches to understand their research challenges better and inch their way forward with their existing hypothesis?

After performing broad searches on their field of specialization, they would then narrow down their focus on their task at hand. How do I get to my next publication? How can I prove that my protein is being expressed? Is it being degraded by proteases? What kind of data will I need to prove my hypothesis? Really, completing a PhD is all about answering questions and pursuing knowledge — ie performing Biology and Biochemistry ReSearch.  But, the tools that are typically used in this search are not efficient or complete. Academic research, even though it is considered at the edge of knowledge and understanding, moves at a snail’s pace because it lacks the right tools.


Improving research efficiency in Biology by using free tools like SciGine

This was the reason that SciGine was created. By using the SciGine search engine, I was hoping that future generations of Biologists and Biochemists would be able to more efficiently tackle their research. The website is set up such that any one can use it by going to SciGine and typing in the search box. The next page shows up results similar to how PubMed and Google organize results as individual biology methods with their search query highlighted along with the author name and time/year of publication. Clicking on an individual method then takes the user to a split screen view consisting of the method steps on the left and a notes section on the right.

I realized that many times, the methods we use for research are amalgamations from multiple articles. So why not design a user interface that allows visitors to take notes while viewing multiple methods easily? On a typical method such as Western Blotting for Kidney Amino-Acid Oxidase this makes method development very simple. I can search for multiple western blot methods using the Browse or Search functionality and then take notes on what exactly I want to do with a protein of interest to me on he right side of the screen. With over 1000+ results from a simple “Western Blot” search, I’m amazed at how much this would have helped me during my PhD.

Saving Biology Methods online for ease-of-access later in lab

Beyond simply taking notes, users can save their methods and protocols and keep a running list of them as part of their Online Methods Notebook. This page is for editing their user profile but also can be used as an online lab notebook. I can envision users using it to keep their day-to-day methods and results online or just to keep a general repository of the methods that they tried out. It’s especially useful to keep this information online because a user can, for example, share their immunoblot method with a lab mate easily and also view their method in lab via a phone or tablet while performing their experiment. There are also option to keep methods private (un-searchable) or public (searchable by everyone) so that important information doesn’t get into the wrong hands.

The ability to upload complete methods online also has the advantage of making it easy to write publications later on. Too often, in a publication, details of their methods are lacking.  By keeping track of what materials were used (especially their part numbers and vendors), the references that were combined to make a method, and how each material was used step-by-step, writing a “Materials and Methods” section for a publication will be a breeze. I want researchers to be able to determine if a chemical from Sigma Aldrich vs. Fisher Scientific was the culprit when it came to their bad data. SciGine will, I expect, improve the quality of research produced by it’s users.

Eventually, I expect there to be more viewers asking and answering questions specific to the methods that they find on SciGine. However, several other resources exist which can help with questions for now. In particular Research Gate and Protocols Online have active communities on their forums that make it easy to get questions answered.

Thank you for reading about the inception of SciGine and how it can be used to advance your Biology and Biochemistry research. Look forward to more posts in the future!