Experiments

Overview of the 3T3-L1 Adipogenesis Process

The 3T3-L1 cell line is a robust and widely used model for studying adipogenesis. The process involves three key stages:

1. Proliferation/Maintenance: Preadipocytes (which resemble fibroblasts) are cultured until they reach confluence.
2. Differentiation: A specific hormonal cocktail is added to the confluent cells, triggering the genetic cascade that transforms them into adipocytes.
3. Maturation & Staining: The differentiated cells accumulate lipids in intracellular droplets, which can be visualized by the fat-soluble dye, Oil Red O.

Part 1: Culturing and Maintaining 3T3-L1 Preadipocytes

This phase involves the routine culture and expansion of the preadipocyte cell line before starting a differentiation experiment.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Maintenance

A. Complete Growth Medium Preparation:

• To a 500 mL bottle of High Glucose DMEM, add:
  • 50 mL of FBCS (for a final concentration of 10%).
  • 5 mL of Penicillin-Streptomycin (for a final concentration of 1% or 100 U/mL Penicillin, 100 μg/mL Streptomycin).
• Store at 4°C. Warm to 37°C in a water bath before use.

B. Thawing and Plating Cryopreserved Cells:

• Warm the Complete Growth Medium to 37°C.
• Thaw the vial of frozen cells rapidly (<1 minute) in a 37°C water bath.
• Wipe the vial with 70% ethanol and transfer it to a biosafety cabinet.
• Gently transfer the cell suspension into a 15 mL conical tube containing 9 mL of pre-warmed medium.
• Centrifuge at 200 x g for 5 minutes.
• Aspirate the supernatant and gently resuspend the cell pellet in 10-12 mL of fresh medium.
• Transfer the cell suspension to a T-75 flask and place it in the incubator.

C. Subculturing (Passaging) Preadipocytes:

• Crucial Note: Do not allow preadipocytes to become 100% confluent during the maintenance phase, as this can reduce their ability to differentiate later. Passage them when they reach 70-80% confluency.
• Aspirate the old medium from the T-75 flask.
• Wash the cell monolayer once with 5-10 mL of sterile PBS.
• Add 2-3 mL of 0.25% Trypsin-EDTA to the flask, ensuring it covers the entire surface. Incubate for 2-5 minutes at 37°C, until cells round up and detach (check under a microscope).
• Add 6-8 mL of Complete Growth Medium to the flask to neutralize the trypsin.
• Transfer the cell suspension to a 15 mL conical tube and centrifuge at 200 x g for 5 minutes.
• Aspirate the supernatant, resuspend the pellet in fresh medium, and re-plate the cells into new flasks at a split ratio of 1:6 to 1:10.
• Change the medium every 2-3 days.

Part 2: Induction of Adipocyte Differentiation

This protocol begins after you have seeded 3T3-L1 cells in culture plates and grown them to the correct state of confluence.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Differentiation

A. Preparation of Stock Solutions and Media:

• Dexamethasone (1 mM stock): Dissolve in ethanol or DMSO. Store at -20°C.
• IBMX (0.5 M stock): Dissolve in DMSO. Store at -20°C.
• Insulin (1 mg/mL stock): Dissolve in 0.02 M HCl or sterile water. Store at -20°C.

B. Preparation of Differentiation Media (example for 100 mL):

• MDI Initiation Medium (Day 0-2):
  • To 100 mL of DMEM + 10% FBS, add:
    • 10 μL of 1 mM DEX stock (Final: 0.1 μM)
    • 100 μL of 0.5 M IBMX stock (Final: 0.5 mM)
    • 100 μL of 1 mg/mL Insulin stock (Final: 1 μg/mL)
• Insulin Medium (Day 2 onwards):
  • To 100 mL of DMEM + 10% FBS, add:
    • 100 μL of 1 mg/mL Insulin stock (Final: 1 μg/mL)

C. Differentiation Timeline:

• Seeding for Differentiation (2-4 Days Before Day 0): Seed 3T3-L1 preadipocytes into the desired plate format (e.g., 6-well plates). Culture them in Complete Growth Medium until they reach 100% confluence.
• Contact Inhibition (Day -2 to Day 0): Once the cells are 100% confluent, allow them to remain in the incubator for an additional 48 hours. This post-confluent state is critical for initiating the differentiation program.
• Day 0: Induction
  • Aspirate the growth medium.
  • Replace it with MDI Initiation Medium. Return the plate to the incubator.
• Day 2 (or Day 3): Media Change
  • Aspirate the MDI Initiation Medium.
  • Replace it with fresh Insulin Medium.
• Day 4 Onwards: Maturation
  • Aspirate and replace the medium with fresh Insulin Medium every 2 days.
  • Monitor the cells under a microscope. Small, refractile lipid droplets should begin to appear in the cytoplasm around Day 4-5 and will grow larger over time. The process is typically complete by Day 8-12, when cells are large, round, and packed with lipid droplets.

Part 3: Oil Red O Staining for Lipid Droplets

This is an endpoint assay to visualize and confirm the accumulation of lipids in the differentiated adipocytes.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Staining

A. Preparation of Staining Solutions:

• Oil Red O Stock Solution (0.5% w/v): Dissolve 0.5 g of Oil Red O powder in 100 mL of 100% isopropanol. Stir overnight to dissolve completely. This solution is stable at room temperature.
• Oil Red O Working Solution:
  • Mix 6 mL of the Oil Red O Stock Solution with 4 mL of distilled water.
  • Let this solution sit at room temperature for 20 minutes.
  • This is the most important step: Filter the working solution through a 0.22 μm or 0.45 μm syringe filter to remove precipitates. This solution must be prepared fresh and used within 2 hours.

B. Staining Procedure (for a 6-well plate):

• Wash: Carefully aspirate the culture medium. Gently wash the cells twice with PBS. Mature adipocytes detach easily, so be very gentle with all washing steps.
• Fixation: Add 1 mL of 10% Formalin to each well. Incubate at room temperature for 30-60 minutes.
• Wash: Aspirate the formalin. Wash the cells twice with distilled water.
• Dehydration Rinse: Aspirate the water. Add 1 mL of 60% isopropanol to each well and incubate for 5 minutes. This step primes the lipid droplets for staining.
• Staining: Aspirate the isopropanol. Add 1 mL of the filtered Oil Red O working solution to each well, ensuring the cell layer is completely covered. Incubate at room temperature for 20-30 minutes.
• Wash: Aspirate the stain. Wash the cells 3-4 times with distilled water until the excess stain is removed and the water runs clear.
• Imaging: Add PBS or water to the wells to keep the cells hydrated. Visualize under a brightfield microscope.

Overview of the Lactobacillus spp. culture

Lactobacillus rhamnosus, a widely studied probiotic bacterium, can be readily cultured in a laboratory setting with the appropriate ingredients and a defined protocol. This guide outlines the necessary components and step-by-step instructions for the successful cultivation of this facultative anaerobe.

I. Laboratory Ingredients and Products

The cornerstone of culturing Lactobacillus rhamnosus is the use of a specific growth medium that provides the necessary nutrients for its proliferation. The most common and recommended medium is the de Man, Rogosa and Sharpe (MRS) medium.

A. Composition of MRS Medium (per 1 liter)

Note: Pre-mixed MRS broth and MRS agar are commercially available from suppliers like Difco, Oxoid, and MilliporeSigma, which can simplify the media preparation process.

Other Essential Laboratory Supplies:

• Sterile culture tubes or flasks: For growing liquid cultures.
• Sterile petri dishes: For solid agar plates.
• Autoclave: For sterilizing media and equipment.
• Incubator: To maintain the optimal growth temperature.
• Anaerobic jar or chamber (optional but recommended): To create an oxygen-deprived environment. Gas-generating packs (e.g., GasPak™) can be used with anaerobic jars.
• Micropipettes and sterile tips: For accurate liquid handling.
• Inoculating loops or sterile swabs: For transferring the bacterial culture.
• pH meter: To adjust the pH of the medium.
• Spectrophotometer (optional): To measure bacterial growth by optical density (OD).
• Lactobacillus rhamnosus starter culture: This can be obtained from a commercial culture collection (e.g., ATCC - American Type Culture Collection) as a freeze-dried or frozen stock.

II. Step-by-Step Culturing Protocol

This protocol outlines the process from preparing the growth medium to harvesting the bacterial cells.

Step 1: Media Preparation

• Weighing and Dissolving: Accurately weigh all the powdered ingredients for the MRS medium (if not using a pre-mixed powder) and dissolve them in approximately 900 mL of distilled water in a large flask or beaker. If making MRS agar, add the agar at this stage.
• Adding Liquids: Add the polysorbate 80 (Tween 80) and mix thoroughly.
• Adjusting pH: Adjust the pH of the medium to 6.2±0.2 at 25°C using 1M HCl or 1M NaOH. This is a critical step as L. rhamnosus prefers a slightly acidic environment for optimal growth.
• Final Volume: Bring the final volume to 1 liter with distilled water.
• Dispensing: Dispense the medium into appropriate culture tubes or flasks for liquid cultures, or into bottles for agar plates.
• Sterilization: Sterilize the medium by autoclaving at 121°C (15 psi) for 15 minutes. Caution: Do not over-autoclave as it can degrade the components of the medium.
• Pouring Plates (for solid medium): If making MRS agar plates, allow the autoclaved medium to cool to around 45-50°C in a water bath before pouring it into sterile petri dishes. Let the plates solidify at room temperature.

Step 2: Inoculation

• Culture Activation: If starting from a freeze-dried culture, rehydrate the pellet in a small amount of MRS broth as per the supplier’s instructions. If using a frozen stock, thaw it at room temperature or in a 37°C water bath.
• Pre-culture (Starter Culture): Inoculate a small volume of sterile MRS broth (e.g., 5-10 mL) with the activated or thawed L. rhamnosus culture. This is your starter culture.
• Incubation of Starter Culture: Incubate the starter culture at 37°C for 18-24 hours. Growth is typically indicated by visible turbidity (cloudiness) in the broth. For optimal growth, anaerobic or microaerophilic conditions are preferred. This can be achieved by using an anaerobic jar with a gas pack or by filling the culture tube to the top and tightening the cap.
• Main Culture Inoculation: After the starter culture has grown, inoculate a larger volume of fresh, sterile MRS broth or the surface of an MRS agar plate. A typical inoculation volume for liquid culture is 1-2% (v/v) of the starter culture (e.g., 1-2 mL of starter culture into 100 mL of fresh broth). For agar plates, use a sterile inoculating loop to streak the starter culture onto the agar surface.

Step 3: Incubation

• Temperature: Incubate the cultures at the optimal temperature for Lactobacillus rhamnosus, which is typically 37°C. Some strains may have slightly different optimal temperatures, so it is advisable to check the specifications for the particular strain being used.

• Atmosphere: As a facultative anaerobe, L. rhamnosus can grow under both aerobic and anaerobic conditions. However, growth is often enhanced in an anaerobic or microaerophilic environment. For liquid cultures, this can be achieved by using sealed containers or an anaerobic chamber. For plate cultures, an anaerobic jar is recommended.

• Time: The typical incubation time is 24 to 48 hours. The exact duration will depend on the desired cell density. Growth can be monitored by observing the turbidity of the liquid culture or the formation of colonies on the agar plate.

Step 4: Harvesting and Assessment

• Harvesting from Liquid Culture:

  • Once the desired growth is achieved (typically in the stationary phase, after 24-48 hours), the bacterial cells can be harvested by centrifugation.
  • Centrifuge the culture at approximately 5,000 x g for 10-15 minutes at 4°C.
  • Discard the supernatant (the clear liquid). The pellet at the bottom of the tube contains the L. rhamnosus cells.
  • The cell pellet can be washed by resuspending it in a sterile buffer solution (e.g., phosphate-buffered saline - PBS) and repeating the centrifugation step.

• Harvesting from Agar Plates:

  • Colonies can be scraped from the surface of the agar plate using a sterile inoculating loop or a cell scraper.
  • Resuspend the collected colonies in a suitable sterile buffer.

• Assessment of Growth (Optional):

  • Optical Density (OD): The growth of a liquid culture can be monitored by measuring the absorbance of the culture at a wavelength of 600 nm (OD600) using a spectrophotometer. An increase in OD correlates with an increase in bacterial cell number.
  • Viable Cell Count: To determine the number of living bacterial cells, a serial dilution and plate count method on MRS agar can be performed. The results are expressed as colony-forming units per milliliter (CFU/mL).

By following this comprehensive protocol, researchers and laboratory personnel can successfully culture Lactobacillus rhamnosus for a wide range of applications, from basic research to the development of probiotic products.

Isolation of Extracellular Vesicles of Lactobacillus rhamnosus

This protocol employs sequential centrifugation to first remove bacterial cells and then uses an Amicon® Ultra-15 (100 kDa) centrifugal filter to concentrate and wash the EVs from the sterile culture supernatant.

I. Laboratory Ingredients and Products

II. Primary Isolation Device: Amicon® Ultra-15 Centrifugal Filter Unit

• Concentrates molecules >100 kDa (including EVs) while filtering out smaller molecules

• MilliporeSigma Cat. No. UFC910024 or UFC910008

III. Step-by-Step Isolation Protocol

This protocol assumes you have already grown a liquid culture of Lactobacillus rhamnosus (e.g., 50 mL or more) to the stationary phase (typically 24-48 hours). Perform all steps under sterile conditions in a biosafety cabinet and keep samples on ice or at 4° C whenever possible to maintain vesicle integrity.

Phase 1: Preparation of Cell-Free Supernatant

Step 1: Pellet Bacterial Cells

• Pour your L. rhamnosus liquid culture into sterile 50 mL conical centrifuge tubes.
• Centrifuge the culture at 10,000 x g for 20 minutes at 4° C. This high speed ensures the firm pelleting of all bacterial cells.
• Carefully decant the supernatant (the liquid portion) into a new, sterile 50 mL conical tube. Be extremely careful not to disturb the bacterial pellet at the bottom.

Step 2: Sterilization Filtration

This is a critical step to remove any remaining bacteria and larger cellular debris.

• Attach a sterile 0.22 µm syringe filter to a new sterile syringe. The size of the syringe should be appropriate for your volume of supernatant.
• Draw the supernatant from the new tube into the syringe.
• Carefully push the supernatant through the 0.22 µm filter into a final sterile container (e.g., another 50 mL tube or a sterile bottle). This is now your cell-free, sterile supernatant.

Phase 2: EV Isolation with Amicon® Ultra-15 (100 kDa) Filter

Step 3: Prepare the Amicon® Filter Unit

• Unbox a new Amicon® Ultra-15, 100 kDa filter unit. It consists of a filter device and a centrifuge tube. Place the filter device into the provided centrifuge tube.
• (Optional but recommended) Pre-rinse the filter by adding 15 mL of sterile PBS, centrifuging at 4,000 x g for 10 minutes at 4° C, and discarding the flow-through. This helps remove any potential preservatives from the filter membrane.

Step 4: Load the Supernatant and Concentrate

• Add up to 15 mL of your cell-free, sterile supernatant to the filter device of the Amicon® unit.
• Cap the unit and place it in a swinging-bucket rotor in a refrigerated centrifuge. Ensure it is properly balanced with another tube of equal weight.
• Centrifuge at 4,000 x g for 15-30 minutes at 4° C. The exact time will depend on the viscosity of your media. The goal is to reduce the volume in the upper chamber to approximately 200-500 µL. Check the volume periodically to avoid spinning to complete dryness.

Step 5: Wash the Concentrated EVs

This step is essential to remove the concentrated media components (salts, amino acids, sugars) that are smaller than 100 kDa but get trapped in the concentrated volume.

• Remove the Amicon® unit from the centrifuge. Discard the flow-through from the centrifuge tube.
• Add 14-15 mL of cold, sterile PBS to the filter device containing your concentrated sample.
• Gently pipette up and down a few times (without touching the membrane) to wash the EVs.
• Cap the unit and centrifuge again at 4,000 x g for 15-25 minutes at 4° C, until the volume is once again reduced to 200-500 µL.
• Repeat this washing step (Step 5.1 - 5.4) at least one more time for higher purity. Two washes are generally recommended.

Step 6: Harvest the Purified EVs

• After the final wash and centrifugation, carefully remove the unit from the centrifuge. Discard the flow- through.
• Place the filter device in a new, clean tube or hold it steady.
• Insert a micropipette with a sterile tip into the filter device and carefully pipette the concentrated EV suspension (the retentate) up and down to rinse the walls and membrane.
• Transfer the final, concentrated EV suspension (typically 200-500 µL) into a sterile 1.5 mL microcentrifuge tube.

IV. Storage and Downstream Analysis

Step 7: Storage

• Label the microcentrifuge tube clearly.
• For short-term storage (1-2 weeks), store the EVs at 4° C.
• For long-term storage, aliquot the sample into smaller volumes (to avoid freeze-thaw cycles) and store at - 80° C.

Important Note on Verification: This protocol isolates particles larger than 100 kDa. To confirm the presence, size, and concentration of EVs in your final sample, further characterization is necessary using techniques such as:

• Nanoparticle Tracking Analysis (NTA): To determine the size distribution and concentration of vesicles.
• Transmission Electron Microscopy (TEM): To visualize the morphology (cup-shaped or spherical) of the vesicles.
• Protein Quantification: Using a BCA or Bradford assay to determine the total protein content of the EV sample.
• Western Blotting: To detect specific protein markers associated with bacterial EVs (if known for L. rhamnosus).

Overview of the RNA workflow

1. Total RNA Extraction: Using TRIzol reagent to lyse cells and purify total RNA, separating it from DNA and proteins.
2. cDNA Synthesis: Using the GeneDepot RT-PreMix Kit to convert the extracted RNA into more stable complementary DNA (cDNA).
3. qPCR Analysis: Using the synthesized cDNA as a template to quantify the relative expression levels of your target genes (Cebpa, Pparg) normalized against a housekeeping gene (Actb).

Part 1: Total RNA Extraction using TRIzol™ Reagent

This protocol is for extracting RNA from cells cultured in a 6-well plate. All steps should be performed in a biosafety cabinet or fume hood, and an RNase-free environment must be maintained throughout to prevent RNA degradation.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for RNA Extraction

1. Cell Lysis (Homogenization)

• Aspirate the culture medium from the 6-well plate.
• Wash the cells once with 1-2 mL of cold, sterile PBS.
• Aspirate the PBS completely.
• Add 1 mL of TRIzol™ Reagent directly to each well.
• Use a cell scraper to scrape the well surface to ensure all cells are lysed in the TRIzol. Pipette the lysate up and down several times to homogenize it.
• Transfer the viscous lysate to a 1.5 mL RNase-free microcentrifuge tube.
• Incubate at room temperature for 5 minutes to permit complete dissociation of nucleoprotein complexes.

2. Phase Separation

• Add 0.2 mL of chloroform to each tube of lysate.
• Securely cap the tube and shake it vigorously by hand for 15 seconds (do not vortex). The solution should become milky.
• Incubate at room temperature for 2-3 minutes.
• Centrifuge the samples at 12,000 x g for 15 minutes at 4°C.
• After centrifugation, the mixture will separate into three phases: a lower red phenol-chloroform phase, a white interphase (containing DNA), and a colorless upper aqueous phase (containing RNA).

3. RNA Precipitation

• Carefully aspirate the upper aqueous phase and transfer it to a new, sterile 1.5 mL RNase-free tube. Be extremely careful not to disturb or transfer any of the white interphase layer.
• Add 0.5 mL of 100% isopropanol to the aqueous phase.
• Invert the tube gently to mix and incubate at room temperature for 10 minutes.
• Centrifuge at 12,000 x g for 10 minutes at 4°C. You should see a small, white gel-like pellet at the bottom of the tube. This is the RNA.

4. RNA Wash

• Carefully decant or aspirate the supernatant, leaving the RNA pellet behind.
• Add 1 mL of 75% ethanol (prepared with RNase-free water).
• Flick the tube to dislodge the pellet and vortex briefly.
• Centrifuge at 7,500 x g for 5 minutes at 4°C.

5. Resuspending RNA

• Carefully aspirate the ethanol wash completely.
• Air-dry the pellet for 5-10 minutes at room temperature. Do not over-dry, as this will make it difficult to dissolve. The pellet should be translucent, not bone-white.
• Resuspend the RNA pellet in 20-50 μL of RNase-free water by pipetting up and down.
• Incubate at 55-60°C for 10 minutes to aid dissolution.

6. Quantification and Quality Check

• Use a NanoDrop spectrophotometer to measure the RNA concentration (A260) and purity (A260/A280 and A260/A230 ratios).
• High-quality RNA will have an A260/A280 ratio of ~2.0 and an A260/A230 ratio of >1.8.
• Store the RNA at -80°C for long-term use.

Part 2: cDNA Synthesis using GeneDepot RT-PreMix Kit

This protocol converts your purified RNA into cDNA, which is a necessary template for qPCR.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for cDNA Synthesis

1. Reaction Setup (on ice)

• The GeneDepot RT-PreMix Kit comes in tubes containing a lyophilized pellet of all necessary reagents (Reverse Transcriptase, dNTPs, primers, buffer, RNase inhibitor).
• Determine the amount of RNA to use. 1 μg of total RNA is a standard starting amount for a 20 μL reaction. Calculate the volume of your RNA sample that contains 1 μg.
• Pipette the 20 μL RNA/water mixture into one of the RT-PreMix tubes.
• Pipette up and down gently to dissolve the pellet completely.

2. Thermal Cycler Program

• Place the reaction tubes in a thermal cycler and run the following program:
  • Reverse Transcription: 45°C for 60 minutes
  • RT Inactivation: 95°C for 5 minutes
• Once the program is complete, place the tubes on ice.

3. Storage

• The resulting cDNA is now ready for qPCR. It can be used immediately or stored at -20°C. For long-term storage, aliquoting is recommended.

qPCR for gene expression

This protocol uses SYBR Green chemistry to quantify the relative abundance of Pparg and Cebpa mRNA, normalized to Actb (beta-actin).

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for qPCR

1. Preparation (on ice)

• Thaw the SYBR Green Master Mix, primer stocks, and cDNA templates on ice.
• Dilute your cDNA template 1:5 or 1:10 with nuclease-free water. A dilution prevents potential inhibitors from the RT reaction from affecting the qPCR.
• Dilute your primer stocks (usually 100 μM) to a working concentration of 10 μM.

2. qPCR Reaction Setup

• It is essential to run each sample in triplicate for each gene. Also include a No-Template Control (NTC) for each primer set (replace cDNA with water) to check for contamination.
• Prepare a master mix for each primer set to ensure consistency across wells.
• For one reaction (example for 10 μL final volume):

Component	Volume	Final Concentration
2x SYBR Green Master Mix	5 μL	1x
Forward Primer (10 μM)	0.5 μL	500 nM
Reverse Primer (10 μM)	0.5 μL	500 nM
Diluted cDNA Template	2 μL	~10-100 ng
Nuclease-Free Water	2 μL	-
Total Volume	10 μL

• Calculate the total volume needed for all your reactions (including triplicates and an extra 10% for pipetting error).
• Aliquot the master mix into the wells of your qPCR plate, then add the respective cDNA template or NTC water.
• Seal the plate firmly with an optical seal and centrifuge briefly to bring all liquid to the bottom of the wells.

3. Real-Time PCR Program

• Place the plate in the qPCR machine and run a standard SYBR Green program:
  • Initial Denaturation: 95°C for 3 minutes (1 cycle)
  • PCR Amplification: 40 cycles of:
    • Denaturation: 95°C for 10 seconds
    • Annealing/Extension: 60°C for 30 seconds (acquire fluorescence data at the end of this step)
  • Melt Curve Analysis: 65°C to 95°C, increasing by 0.5°C per step (1 cycle). This is a critical QC step to verify the specificity of the amplification.

4. Data Analysis (Relative Quantification using the ΔΔCt Method)

• Export the raw Ct (Cycle threshold) values from the qPCR software.
• Average the Ct values for each set of triplicates.
• Step 1: Calculate ΔCt. Normalize the target gene to the housekeeping gene (Actb). ΔCt = Ct(target gene, e.g., Pparg) - Ct(housekeeping gene, Actb)
• Step 2: Calculate ΔΔCt. Normalize the treated/differentiated sample to the untreated/undifferentiated control sample. ΔΔCt = ΔCt(differentiated sample) - ΔCt(undifferentiated control)
• Step 3: Calculate Fold Change. Fold Change = 2^(−ΔΔCt)
• A fold change > 1 indicates upregulation of the gene in the differentiated sample compared to the control. You should see a significant upregulation of Pparg and Cebpa in your successfully differentiated 3T3-L1 cells.

Overview of the Western Blot workflow

Overview of the Workflow

1. Total Protein Extraction & Quantification: Using RIPA buffer to lyse cells and purify total protein, separating it from other cellular components. Protein concentration is then measured using a BCA or Bradford assay.

2. SDS-PAGE: Denaturing the extracted proteins and separating them by molecular weight using polyacrylamide gel electrophoresis.

3. Protein Transfer (Blotting): Transferring the separated proteins from the gel onto a PVDF or nitrocellulose membrane.

4. Immunodetection: Using specific primary and secondary antibodies to detect the target protein on the membrane.

5. Signal Detection and Analysis: Visualizing the protein bands using a chemiluminescent substrate and quantifying their relative expression levels normalized to a loading control (e.g., GAPDH or β-actin).

Part 1: Total Protein Extraction and Quantification

This protocol is for extracting protein from cells cultured in a 6-well plate. All steps should be performed on ice or at 4°C to prevent protein degradation by proteases.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Protein Extraction

1. Cell Lysis

• Aspirate the culture medium from the 6-well plate.
• Wash the cells once with 1-2 mL of ice-cold, sterile PBS.
• Aspirate the PBS completely.
• Add 100-150 μL of ice-cold RIPA buffer (pre-mixed with protease/phosphatase inhibitors) directly to each well.
• Use a cell scraper to scrape the well surface, collecting the cells in the lysis buffer.
• Transfer the lysate to a pre-chilled 1.5 mL microcentrifuge tube.
• Incubate on ice for 15-30 minutes, vortexing briefly every 10 minutes.

2. Lysate Clarification

• Centrifuge the lysate at 14,000 x g for 15 minutes at 4°C. This will pellet cell debris.
• Carefully transfer the clear supernatant, which contains the soluble proteins, to a new pre-chilled 1.5 mL tube. Discard the pellet.

3. Protein Quantification

• Perform a protein quantification assay (e.g., BCA) according to the manufacturer’s protocol to determine the concentration of your protein samples.

4. Storage

• The protein lysate can be used immediately or stored at -80°C for long-term use.

Part 2: SDS-PAGE for Protein Separation

This protocol separates the extracted proteins based on their molecular weight.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for SDS-PAGE

1.Sample Preparation

• Based on the quantification results, calculate the volume of protein lysate needed for 20-30 µg of total protein per lane.
• In a 1.5 mL tube, mix your calculated volume of lysate with 4x Laemmli sample buffer to a final concentration of 1x. Add nuclease-free water to reach the final desired volume (e.g., 20 µL).
• Heat the prepared samples at 95-100°C for 5 minutes to complete denaturation.

2.Gel Electrophoresis

• Assemble the electrophoresis chamber according to the manufacturer’s instructions and fill it with 1x running buffer.
• Load 5-10 µL of the protein ladder into the first well.
• Load your prepared protein samples (e.g., 20 µL) into the subsequent wells.
• Run the gel at a constant voltage (e.g., 120-150 V) for 45-90 minutes, or until the dye front reaches the bottom of the gel.

Part 3: Protein Transfer (Blotting)

This protocol transfers the proteins from the gel to a membrane for antibody detection.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Transfer

1.Preparation

• Cut a piece of PVDF membrane and filter paper to the size of the gel.
• Activate the PVDF membrane by soaking it in 100% methanol for 1 minute, followed by a brief rinse in deionized water, and then equilibration in 1x Transfer Buffer for at least 5 minutes.
• Equilibrate the gel and filter papers in 1x Transfer Buffer for 5-10 minutes.

2.Assemble the Transfer Stack

• Assemble the transfer “sandwich” according to the manufacturer’s instructions (the order is critical and depends on the system). A typical order for semi-dry transfer is: Anode plate -> presoaked filter paper -> PVDF membrane -> Gel -> pre-soaked filter paper -> Cathode plate.
• Use a roller to remove any air bubbles between the layers.

3.Transfer

• Place the transfer stack in the blotting apparatus and run the transfer program (e.g., 25V for 30 minutes for a standard semi-dry transfer).

4.Confirmation (Optional but recommended)

• After transfer, you can briefly stain the membrane with Ponceau S solution to visualize total protein and confirm transfer efficiency. Wash off the stain completely with water or TBST before blocking.

Part 4: Immunodetection

This protocol uses antibodies to detect a specific protein of interest.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Immunodetection

1.Blocking

• Place the membrane in a small container and add enough blocking buffer (e.g., 5% milk in TBST) to fully submerge it.
• Incubate for 1 hour at room temperature on a shaker.

2.Primary Antibody Incubation

• Dilute the primary antibody in fresh blocking buffer to the concentration recommended by the manufacturer.
• Pour off the blocking solution and add the diluted primary antibody solution to the membrane.
• Incubate overnight at 4°C on a shaker (or for 1-2 hours at room temperature).

3.Washing

• Pour off the primary antibody solution. Wash the membrane 3 times for 5-10 minutes each with TBST on a shaker.

4.Secondary Antibody Incubation

• Dilute the HRP-conjugated secondary antibody in fresh blocking buffer.
• Add the diluted secondary antibody solution to the membrane.
• Incubate for 1 hour at room temperature on a shaker.

5.Final Washes

• Pour off the secondary antibody solution. Wash the membrane 3 times for 10 minutes each with TBST on a shaker.

6.Signal Development

• Prepare the ECL substrate by mixing the two components according to the manufacturer’s instructions.
• Place the membrane, protein-side up, on a clean surface. Pipette the ECL substrate evenly over the surface of the membrane.
• Incubate for 1-5 minutes. Do not let the membrane dry out.

Part 5: Signal Detection and Data Analysis

This protocol captures the chemiluminescent signal and quantifies the results.

I. Laboratory Ingredients and Products

2. Step-by-Step Protocol for Analysis

1.Image Acquisition

• Carefully place the membrane inside the imaging system, ensuring there are no wrinkles.
• Capture the image. Use an auto-exposure setting or take multiple manual exposures to ensure the signal is strong but the bands are not saturated (over-exposed).

2.Data Analysis

• Use image analysis software (e.g., ImageJ) to measure the integrated density of each band.
• Normalization: To correct for unequal sample loading, normalize the density of your target protein band to the density of a loading control band (e.g., GAPDH, β-actin) from the same lane.
• Relative Quantification: The final results are expressed as the relative fold change in protein expression compared to a control sample.

Overview of the Cloning Workflow

Overview of the Workflow

1. PCR Amplification of Insert DNA: The gene of interest (the “insert”) is amplified from a template (e.g., cDNA, gDNA) using PCR. The primers are designed to include restriction enzyme recognition sites.

2. Vector and Insert Preparation (Digestion & Purification): The amplified insert and the plasmid vector are digested with the same restriction enzymes to create compatible ends (sticky or blunt). The desired DNA fragments are then purified.

3. Ligation: The digested vector and insert are joined into a single, circular recombinant plasmid using the T4 DNA Ligase enzyme.

4. Transformation: The completed recombinant plasmid is introduced into a host cell, typically competent E. coli.

5. Selection & Screening: Colonies of bacteria that successfully incorporated a plasmid are selected using antibiotic resistance. These colonies are then screened to identify those containing the correct insert.

6. Plasmid Miniprep and Verification: A selected colony is grown in a liquid culture, and the plasmid DNA is extracted (Miniprep). The success of the cloning is confirmed by final verification, typically via Sanger sequencing.

Part 1: PCR Amplification of Insert DNA

This step amplifies the target gene fragment for cloning

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2. Step-by-Step Protocol for PCR

1. PCR Reaction Setup (on ice)

• Prepare the PCR reaction mix in a sterile PCR tube as shown in the table below.

Component	Volume (for 50 μL)	Final Concentration
5x HF Buffer	10 μL	1x
10 mM dNTPs	1 μL	200 μM
Forward Primer (10 μM)	2.5 μL	0.5 μM
Reverse Primer (10 μM)	2.5 μL	0.5 μM
Template DNA (1-10 ng/μL)	1 μL	1-10 ng
High-fidelity DNA Polymerase	0.5 μL	1 unit
Nuclease-Free Water	Up to 50 μL

2. Run Thermal Cycler Program

• Place the PCR tubes in a thermal cycler and run a standard program. (Adjust annealing temperature based on primers).

Step	Temperature	Time	Cycles
Initial Denaturation	98°C	30 sec	1
Denaturation	98°C	10 sec
Annealing	55-65°C	30 sec
Extension	72°C	30-60 sec / kb
Final Extension	72°C	5-10 min	1
Hold	4°C	∞	1

3. Confirm Amplification

• Run a small amount (e.g., 5 μL) of the PCR product on an agarose gel to verify that a DNA band of the expected size has been amplified.

Part 2: Vector and Insert Preparation (Digestion & Purification)

This step prepares the PCR product (insert) and plasmid vector by cutting them with restriction enzymes.

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2. Step-by-Step Protocol for Preparation

1. Restriction Digest

• Set up separate digestion reactions for the vector and the purified PCR product using the same restriction enzymes.

Component	Vector (e.g.)	Insert (e.g.)
Plasmid Vector DNA (e.g., 1 μg)	X μL	-
Purified PCR Product (e.g., 500 ng)	-	Y μL
10x Reaction Buffer	3 μL	5 μL
Restriction Enzyme 1 (e.g., EcoRI)	1 μL	1 μL
Restriction Enzyme 2 (e.g., XhoI)	1 μL	1 μL
Nuclease-Free Water	Up to 30 μL	Up to 50 μL

• Mix well and incubate at the enzyme’s optimal temperature (usually 37°C) for 1-2 hours.

2. Agarose Gel Electrophoresis

• Load the digested DNA onto an agarose gel to separate the fragments. Confirm that the vector is linearized and the insert is the correct size.

3. Gel Extraction

• Using a UV transilluminator, locate and excise the desired DNA bands (linearized vector, insert) from the gel with a clean scalpel.
• Use a Gel Extraction Kit to purify the DNA from the agarose slice according to the manufacturer’s protocol.

4. Quantification

• Measure the concentration of the purified vector and insert DNA using a spectrophotometer (e.g., NanoDrop).

Part 3: Ligation (Joining Insert and Vector)

This step uses T4 DNA Ligase to join the prepared vector and insert into a single circular plasmid.

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2. Step-by-Step Protocol for Ligation

1. Ligation Reaction Setup

• For optimal efficiency, set up the reaction with a molar ratio of insert to vector of 3:1 or 5:1. (Online calculators are available for this).

Component	Volume
Linearized Vector (e.g., 50 ng)	X μL
Insert (3:1 molar ratio)	Y μL
10x T4 DNA Ligase Buffer	1 μL
T4 DNA Ligase	0.5 μL
Nuclease-Free Water	Up to 10 μL

2. Incubation

• Mix the components gently and incubate at room temperature (20-25°C) for 1 hour or at 16°C overnight.

Part 4: Transformation

This step introduces the ligation product into competent E. coli

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2. Step-by-Step Protocol for Transformation

1.Add DNA

• Thaw a tube of competent cells (50 µL) on ice. Add 5-10 µL of the ligation reaction product and mix gently with a pipette tip.
• Incubate on ice for 20-30 minutes.

2.Heat Shock

• Place the tube in a 42°C water bath for exactly 45-60 seconds.
• Immediately transfer the tube back to ice for 2 minutes.

3.Recovery

• Add 250 µL of SOC medium and incubate at 37°C for 1 hour with shaking (200 rpm).

Part 5: Selection, Miniprep, and Verification

This final stage involves selecting transformed bacteria and verifying the cloned plasmid.

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2. Step-by-Step Protocol for Verification

1. Plating and Selection

• Spread ~100 μL of the recovered cell culture onto an LB agar plate containing the appropriate antibiotic.
• Incubate the plate overnight at 37°C until single colonies appear.

2. Liquid Culture and Miniprep

• Select 3-5 individual colonies and inoculate each into 3-5 mL of LB broth containing the appropriate antibiotic.
• Grow the cultures overnight at 37°C in a shaking incubator (200 rpm).
• Pellet the bacteria by centrifugation and use a Plasmid Miniprep Kit to extract the plasmid DNA from each culture.

3. Final Verification

• Restriction Digest: Perform a diagnostic digest on the purified plasmid DNA using the same restriction enzymes from the cloning step. Run on a gel to confirm the presence of the vector and insert bands.
• Sanger Sequencing: For absolute confirmation, send the purified plasmid DNA to a commercial service for sequencing. This will verify the exact sequence of the insert, its orientation, and that no mutations were introduced.

4. Storage

• Store the confirmed plasmid DNA at -20°C. For long-term storage, create a glycerol stock of the verified bacterial clone.

Overview of the Protein Expression

Overview of the Workflow

1.Transformation into Expression Host: The pET-28b expression vector is transformed into a suitable E. coli expression strain, typically BL21(DE3), which contains the T7 RNA polymerase gene required for expression.

2.Small-Scale Expression Trial: A small pilot culture is performed to optimize expression conditions, such as IPTG concentration, induction temperature, and time, to maximize the yield of soluble protein.

3.Large-Scale Culture and Induction: Based on the optimized conditions, a large-scale culture is grown and protein expression is induced with IPTG.

4.Cell Harvest and Lysis: The bacterial cells are harvested by centrifugation and then lysed, typically by sonication, to release the expressed protein.

5.Analysis of Expression and Solubility: The total cell lysate, as well as the soluble and insoluble fractions, are analyzed by SDS-PAGE to confirm the expression of the target protein and determine its solubility.

Part 1: Transformation into BL21(DE3) Expression Host

This protocol moves your pET-28b plasmid into the E. coli strain that will be used for protein production.

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2. Step-by-Step Protocol for Transformation

1. Thaw one vial (50 μL) of BL21(DE3) competent cells on ice.

2. Add 1-2 μL (approx. 10-50 ng) of your pET-28b plasmid DNA to the cells. Swirl gently to mix.

3. Incubate the tube on ice for 30 minutes.

4. Perform a heat shock by placing the tube in a 42°C water bath for exactly 45 seconds.

5. Immediately transfer the tube back to ice for 2 minutes.

6. Add 250 μL of SOC medium to the tube and incubate at 37°C for 1 hour in a shaking incubator.

7. Spread 100 μL of the cell culture onto a pre-warmed LB agar plate containing 50 μg/mL Kanamycin.

8. Incubate the plate overnight at 37°C.

Part 2: Small-Scale Expression Trial (Optimization)

This pilot experiment is critical to identify the best conditions for expressing your protein in a soluble form.

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2. Step-by-Step Protocol for the Trial

1. Inoculate a single colony from your transformation plate into 5 mL of LB + Kanamycin. Grow overnight at 37°C with shaking. This is your starter culture.

2. The next morning, inoculate 50 mL of LB + Kanamycin with 500 μL of the overnight starter culture (a 1:100 dilution).

3. Grow the culture at 37°C with vigorous shaking until the optical density at 600 nm (OD₆₀₀) reaches 0.5–0.8 (mid-log phase).

4. Remove a 1 mL aliquot of the un-induced culture. Centrifuge, discard the supernatant, and freeze the cell pellet. This is your “Pre-Induction” sample.

5. Divide the remaining culture into smaller, equal volumes (e.g., 4 x 10 mL).

6. Induce each sub-culture with a different final concentration of IPTG (e.g., 0.1 mM, 0.5 mM, 1.0 mM). Leave one un-induced as a negative control.

7. Incubate the cultures under different temperature conditions to test for solubility (e.g., two at 37°C for 4 hours, two at 18°C overnight).

8. After the induction period, measure the final OD₆₀₀, then harvest 1 mL from each culture by centrifugation. Store the pellets at -20°C for analysis in Part 5.

Part 3: Large-Scale Culture and Induction

Using the optimal conditions identified in Part 2, you now scale up the culture for protein production.

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2. Step-by-Step Protocol for Large-Scale Growth

1. Inoculate 10-20 mL of LB + Kanamycin with a single colony and grow overnight at 37°C.

2. The next morning, inoculate 1 L of LB + Kanamycin in a 2 L baffled flask with the entire overnight starter culture.

3. Grow at 37°C with vigorous shaking (approx. 220 rpm). Monitor the OD₆₀₀ every hour.

4. When the OD₆₀₀ reaches 0.6–0.8, remove a 1 mL “Pre-Induction” sample.

5. Induce the culture by adding IPTG to the optimal final concentration determined in Part 2.

6. Reduce the incubator temperature and continue shaking for the optimal time determined in Part 2 (e.g., 18°C for 16-20 hours).

Part 4: Cell Harvest and Lysis by Sonication

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2. Step-by-Step Protocol for Lysis

1. Harvest Cells: Transfer the induced culture to large centrifuge bottles. Centrifuge at 6,000 x g for 15 minutes at 4°C. Carefully discard the supernatant.

2. Resuspend Pellet: Resuspend the cell pellet in 20-30 mL of ice-cold Lysis Buffer.

3. Enzymatic Lysis: Add Lysozyme (to 1 mg/mL), DNase I, and a protease inhibitor cocktail. Incubate on ice for 30 minutes with gentle rocking.

4. Mechanical Lysis (Sonication): Place the tube containing the resuspension in an ice-water bath. Immerse the sonicator probe into the suspension. Sonicate with short pulses (e.g., 10 seconds ON, 20 seconds OFF) for a total of 5-10 minutes of “ON time. The solution should become less viscous.

5. Clarify Lysate: Centrifuge the lysate at high speed (>15,000 x g) for 30 minutes at 4°C.

6. Collect Fractions: Carefully collect the supernatant (the soluble fraction) into a new, clean tube. The pellet is the insoluble fraction. Store both at -20°C or proceed directly to purification/analysis.

Part 5: Analysis of Protein Expression and Solubility

This final step uses SDS-PAGE to confirm that your target protein was expressed and to determine if it is in the soluble or insoluble fraction.

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2. Step-by-Step Protocol for Analysis

• Prepare Samples for SDS-PAGE:

  • Pre- and Post-Induction: Resuspend the “Pre-Induction” and “Post-Induction” cell pellets (from Part 3) in 100 μL of 1x Laemmli buffer.

  • Soluble Fraction: Mix 20 μL of the soluble supernatant (from Part 4) with 20 μL of 2x Laemmli buffer.

  • Insoluble Fraction: Resuspend the insoluble pellet (from Part 4) in 200 μL of lysis buffer, then mix 20 μL of this suspension with 20 μL of 2x Laemmli buffer.

  • Boil all prepared samples at 95-100°C for 5-10 minutes.

• SDS-PAGE: Load 10-15 μL of each sample, along with a protein ladder, onto a polyacrylamide gel. Run the gel until the dye front reaches the bottom.

• Staining: Stain the gel with Coomassie Blue for 1 hour, then destain until the background is clear and protein bands are sharp.

• Analyze the Gel:

  • Compare the “Pre-Induction” and “Post-Induction” lanes. Successful expression is indicated by a prominent new band at the expected molecular weight of your protein in the post-induction lane.

  • Compare the “Soluble” and “Insoluble” lanes. If the new protein band is primarily in the soluble lane, the protein is soluble and ready for purification. If it is primarily in the insoluble lane, it has formed inclusion bodies, and purification will require a denaturing/refolding strategy.