1. Prepare the Wash Solutions: Add 21 mL of ACS grade 100% ethanol to the bottle labeled miRNA Wash Solution 1. Mix well. Place a check mark in the empty box on the label to indicate that the ethanol has been added. Add 40 mL of ACS grade 100% ethanol to the bottle labeled Wash Solution 2/3. Mix well. Place a check mark in the empty box on the label to indicate that the ethanol has been added.
2. Tissue Disruption: Quickly cut the tissue into pieces small enough for either storage or disruption. Weigh the tissue sample (for samples to be stored in RNAlater, this can be done later); Inactivate RNases by one of the following methods:
a. Drop the sample into RNAlater-tissue must be cut to 0.5 cm in at least one dimension for good penetration of the RNAlater.
b. Disrupt the sample in Lysis/Binding Buffer.
c. Freeze the sample in liquid nitrogen-tissue pieces must be small enough to freeze in a few seconds. When the liquid nitrogen stops churning, it indicates that the tissue is completely frozen. Once frozen, remove the tissue from the liquid nitrogen and store it in an airtight container at -70C or colder.
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Measure or estimate the weight of the sample; Place 10 volumes of Lysis/Binding Buffer per tissue mass into a plastic weigh boat or tube on ice; Grind frozen tissue to a powder with liquid nitrogen in a prechilled mortar and pestle sitting in a bed of dry ice; Using a prechilled metal spatula, scrape the powdered tissue into the Lysis/Binding Buffer, and mix rapidly.
3. Organic Extraction: Add 1/10 volume of miRNA Homogenate Additive to the cell or tissue lysate (or homogenate), and mix well by vortexing or inverting the tube several times; Leave the mixture on ice for 10 min; Add a volume of Acid-Phenol:Chloroform that is equal to the Lysate volume before addition of the miRNA Homogenate Additive; Vortex for 30-60 sec to mix; Centrifuge for 5 min at maximum speed (10,000 g) at room temperature to separate the aqueous and organic phases. After centrifugation, the interphase should be compact; if it is not, repeat the centrifugation; Carefully remove the aqueous (upper) phase without disturbing the lower phase, and transfer it to a fresh tube. Note the volume remove.
4. Total RNA Isolation: Preheat Elution Solution or nuclease-free water to 95C for use in eluting the RNA from the filter at the end of the procedure; Add 1.25 volumes of room temperature 100% ethanol to the aqueous phase, and mix thoroughly; Pipet the lysate/ethanol mixture (from the previous step) onto the Filter Cartridge. Up to 700 L can be applied to a Filter Cartridge at a time; Centrifuge for ~15 sec to pass the mixture through the filter; Discard the flow-through, and repeat until all of the lysate/ethanol mixture is through the filter. Reuse the Collection Tube for the washing steps; Apply 700 L miRNA Wash Solution 1 (working solution mixed with ethanol) to the Filter Cartridge and centrifuge for ~5C10 sec or use a vacuum to pull the solution through the filter. Discard the flow-through from the Collection Tube, and replace the Filter Cartridge into the same Collection Tube; Apply 500 L Wash Solution 2/3 (working solution mixed with ethanol) and draw it through the Filter Cartridge as in the previous step; Repeat with a second 500 L aliquot of Wash Solution 2/3; After discarding the flow-through from the last wash, replace the Filter Cartridge in the same Collection Tube and spin the assembly for 1 min to remove residual fluid from the filter; Transfer the Filter Cartridge into a fresh Collection Tube. Apply 100 L of pre-heated (95C) Elution Solution or nuclease-free water to the center of the filter, and close the cap. Spin for ~20C30 sec at maximum speed to recover the RNA; Collect the eluate (which contains the RNA) and store it at C20C.
5. Enrichment Procedure for Small RNAs: Add 1/3 volume 100% ethanol, and mix thoroughly; Add 2/3 volume 100% ethanol and mix thoroughly; Pass the mixture through a second Filter Cartridge, and discard the flow-through; Wash the filter with 700 L miRNA Wash Solution 1; Wash the filter twice with 500 L Wash Solution 2/3; Elute RNA with 100 L 95C Elution Solution or Nuclease-free Water; Collect the eluate (which contains the RNA) and store it at C20C.
Always on Time
Marked to Standard
2.2.2 MicroRNA Microarray
1. Total RNA (100 ng) + Labeling Spike-In (optional): Phosphatase Treatment, incubate 30 minutes, 37C
2. Dephosphorylated RNA*: Add DMSO; Heat, ice; Assemble Labeling Reaction, incubate 2 hours, 16C
3. Labeled RNA*: Desalt with Spin Column
4. Desalted Labeled RNA*: Dry sample with vacuum concentrator, approximately 1 hour, 45C to 55C; Assemble Hybridization Mixture + Hyb Spike-In (optional); Heat, ice Hybridize 20 hours, 55C, 20 RPM; Wash, scan
5. miRNA Profile
*: The sample can be stored frozen at -80C if needed.
1. Prepare the labeling reaction: Dilute total RNA sample to 25 ng/L in 1 TE pH 7.5 or DNase/RNase-free water; Add 4 L (100ng) of the diluted total RNA to a 1.5 mL microcentrifuge tube and maintain on ice; Immediately prior to use, prepare the Calf Intestinal Alkaline Phosphatase (CIP) Master Mix; Add 3 L of the CIP Master Mix to each sample tube for a total reaction volume of 7 L. Gently mix by pipetting; Dephosphorylate the sample by incubating the reaction at 37C in a circulating water bath or heat block for 30 minutes.Add 5 L of 100% DMSO to each sample; Incubate samples at 100C in a circulating water bath or heat block for 5 to 10 minutes; Immediately transfer to ice-water bath; Warm the 10 T4 RNA Ligase Buffer at 37C and vortex until all precipitate is dissolved; Immediately prior to use, prepare the Ligation Master Mix by gently mixing the components; Immediately add 8.0 L of the Ligation Master Mix to each sample tube for a total reaction volume of 20 L; Gently mix by pipetting and gently spin down; Incubate at 16C in a circulating water bath or cool block for 2 hours.
2. Sample purification and hybridization: Add 30 L of RNase-Free Water or 1 TE pH 7.5 to the labeled sample for a total volume of 50 L; Without disturbing the gel bed, pipette the 50 L sample onto the gel bed from step 6 above; Elute the purified sample by spinning the microcentrifuge tubes containing the columns for 4 minutes at 1000 g in a centrifuge; Discard the columns and keep the miRNA sample-containing flow-through on ice; Check that the final flow-through is translucent and slightly pink. The flow-through volume needs to be uniform across the samples and close to 50 L; After sample purification, completely dry the samples. Use a vacuum concentrator at 45 to 55C or on the medium-high heat setting. This step may take up to 1 hour after column purification; Resuspend the dried sample in 17 L of nuclease-free water when the Hyb Spike-In solution is used and 18 L when the Hyb Spike-In solution is not used; 4.5 L of the 10 GE Blocking Agent to each sample; Add 22.5 L of 2 Hi-RPM Hybridization Buffer to each sample for a total of 45 L.Mix well but gently on a vortex mixer; Incubate at 100C for 5 minutes; Immediately transfer to an ice water bath for 5 minutes.
3. hybridization assembly, microarray washing and scanning: Load a clean gasket slide into the Agilent SureHyb chamber base with the label facing up and aligned with the rectangular section of the chamber base; Slowly dispense all of the volume of the hybridization sample onto the gasket well in a drag and dispense manner; Slowly place an array active side down onto the SureHyb gasket slide, so that the Agilent-labeled barcode is facing down and the numeric barcode is facing up. Verify that the sandwich-pair is properly aligned; Place the SureHyb chamber cover onto the sandwiched slides and slide the clamp assembly onto both pieces; Hand-tighten the clamp onto the chamber; Vertically rotate the assembled chamber to wet the gasket and assess the mobility of the bubbles. If necessary, tap the assembly on a hard surface to move stationary bubbles; Place assembled slide chamber in rotisserie in a hybridization oven set to 55C. Set your hybridization rotator to rotate at 20 rpm; Hybridize at 55C for 20 hours; Wash the microarray slides with solutions; Assemble the slides into a version B slide holder; Place assembled slide holders into either a B or C scanner carousel; Verify scan settings for miRNA; Extract data using Agilent Feature Extraction Software (FE version 10.7).
2.2.3 Revere Transcription
Reverse Transcription is carried out with the SuperScript First-Strand Synthesis System for RT-PCR. The following procedure is based on Invitrogens protocol.
4. Prepare the following RNA/primer mixture in each tube:
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Total RNA 5 ?g
random hexamers (50 ng/?l) 3 ?l
10 mM dNTP mix 1 ?l
DEPC H2O to 10 ?l
5. Incubate the samples at 65?C for 5 min and then on ice for at least 1 min.
6. Prepare reaction master mixture. For each reaction:
10 RT buffer 2 ?l
25 mM MgCl2 4 ?l
0.1 M DTT 2 ?l
RNaseOUT 1 ?l
7. Add the reaction mixture to the RNA/primer mixture, mix briefly, and then place at room temperature for 2 min.
8. Add 1 ?l (50 units) of SuperScript II RT to each tube, mix and incubate at 25?C for 10 min.
9. Incubate the tubes at 42?C for 50 min, heat inactivate at 70?C for 15 min, and then chill on ice.
10. Add 1 ?l RNase H and incubate at 37?C for 20 min.
11. Store the 1st strand cDNA at -20?C until use for real-time PCR.
2.2.4 Real-time PCR
1. Normalize the primer concentrations and mix gene-specific forward and reverse primer pair. Each primer (forward or reverse) concentration in the mixture is 5 pmol/?l.
2. A real-time PCR reaction mixture can be either 50 ?l or 25 ?l. Prepare the following mixture in each optical tube.
25 ?l SYBR Green Mix (2x)
0.5 ?l liver cDNA
2 ?l primer pair mix (5 pmol/?l per primer)
22.5 ?l H2O OR 12.5 ?l SYBR Green Mix (2x)
0.2 ?l liver cDNA
1 ?l primer pair mix (5 pmol/?l per primer)
11.3 ?l H2O
3. Set up the experiment and the following PCR program on ABI Prism SDS 7700.
4. After PCR is finished, remove the tubes from the machine. The PCR specificity is examined by 3% agarose gel using 5 ?l from each reaction.
5. Put the tubes back in SDS 7700 and perform dissociation curve analysis with the saved copy of the setup file.
6. Analyze the real-time PCR result with the SDS 7700 software. Check to see if there is any bimodal dissociation curve or abnormal amplification plot.
2.2.5 Cell Counting
1. Place cell counting chamber on a fresh Kimwipe.
2. Label slide chamber #1 and chamber #2 on the white margin of the chamber. Take care to make sure the clear portion of the counting chamber are not touched.
3. Prepare cells by inverting cell containing tube 10 times, pipetting up and down 10 times. Do not shake! Shaking sample will generate bubbles.
4. Stain cells with trypan blue when viable cell count is required. Take 50 l or more cell suspension for mixing with trypan blue.
5. Take 20 l cell suspension after further mixing to load chamber #1. Immediately take another 20 l cell suspension and load chamber #2.
6. Insert the loaded chamber into Cellometer Auto sample slot, and gently push the slide to the stop. The instrument will count and analyze only the chamber inside the system. To measure the second chamber, remove the slide from Cellometer Auto, turn around the slide, and insert the second chamber in the instrument.
7. Click "Count" button while the cell image is displayed.
2.2.6 Accurate Cell Counting for Cells in 6-Well Plate
1. Weight for each well a small falcon
2. Put 2 ml of pre-warmed RPMI+++ medium in a small falcon
3. Aspire the media from the cells
4. Wash the cells with 2 ml pre-warmed PBS
5. Add 1 ml pre-warmed accutase to the cells
6. Detach the cells by pipetting 10-15 times the accutase on the cells
7. Transfer the cell accutase suspension to the small falcon (Step1).
8. Rinse the well with 2 ml pre-warmed RPMI+++-medium
9. Centrifuge the falcon at 1440 rpm for 4 min at room temperature
10. Carefully remove the supernatant of the pellet with a pipett
11. Weight the final volume of each falcon containing cells and media and determine the final volume in each falcon.
12. Resuspend the pellet by pipetting 10-15 times
13. Take a 100 l aliquot of the cell suspension to determine the cell number 4 times.
2.2.7 Cell Lysate Extracts
1. Put the 6-well plate on ice
2. Remove the old medium and wash adherent cells twice in the dish or flask with ice-cold PBS and drain off PBS.
3. Add ice-cold modified RIPA buffer to cells (1 ml per 107 cells/100 mm dish/150 cm2 flask; 0.5 mL per 5 106 cells/60 mm dish/75 cm2 flask).
4. Note: This Cell Extraction Buffer must be supplemented with Phosphatase inhibitor Cocktail tablet and Protease Inhibitor Cocktail tablet just prior to use to make Complete Cell Extraction Buffer. Addition of the Protease Inhibitor Cocktail is necessary to inhibit proteolysis in cell extracts and Phosphatase inhibitor Cocktail preserves the phosphorylation state of proteins during and after cell lysis or tissue protein extraction
5. Put on ice for 10 min and -80C for 20 min
6. Scrape adherent cells off the dish or flask with either a rubber policeman or a plastic cell scraper that has been cooled in ice-cold distilled water. Transfer the cell suspension into a centrifuge tube.
7. Treat the lysate with ultrasound, which breaks the cell walls and shears the DNA into sizes that will not affect the viscosity of the samples.
8. Centrifuge the lysate at 14,000 g in a pre-cooled centrifuge for 15 minutes. Immediately transfer the supernatant to a fresh centrifuge tube and discard the pellet.
2.2.8 Pierce Assay
1. Dilute the contents of one Albumin Standard sample into several clean vials, preferably using the same diluent such as RIPA.
Vial Volume of Diluent (L) Albumin Standard (L)
0 100 0 = Blank
7 100 100
6 100 100 of vial 7
5 100 100 of vial 6
4 100 100 of vial 5
3 100 100 of vial 4
2 100 100 of vial 3
1 100 100 of vial 2
2. Prepare WR by mixing 50 parts of Pierce BCA (bicinchoninic acid) Reagent A with 1 part of BCA Reagent B (50:1, Reagent A:B).
1. Pipette 25L of each standard or unknown sample replicate into a microplate well (working range = 20-2000g/mL).
2. Add 200L of the WR to each well and mix plate thoroughly on a plate shaker for 30 seconds.
3. Cover plate and incubate at 37C for 30 minutes.
4. Cool plate to room temperature. Measure the absorbance at or near 562nm on a plate reader.
2.2.9 Western Blot
1. Precasted gels from Invitrogen 4/20 %: max volume to be added in each well 40l; clamp two gels, add buffer x in the middle to check if does not leak; wash each well with buffer by syringe to avoid air bubble and change buffer in well; add 6 l of marker to first and last well; load 10 or 20 g of protein depending on Ab; run at max mA and 125 V for 120 min.
2. Blotting: prepare a blotting box in ice; open carefully the cast; rinse the gels in blotting buffer (Blotting buffer: 100 ml 10 buffer; 700 ml Aqua dest; 200 ml methanol); incubate the membrane in methanol for 10 sec; then wash it in water; prepare the gel for transfer and watch out it should not run dry; sponge, filter paper, gel, membrane, filter paper, sponge; remove air bubble (roll with falcon), close the cage and put into blotting box; match the migration direction (black to black); run at 400 mA and max Volt for 70 min; wash the blots 1 TBST, 1 methanol and let it dry in filter paper for 30 min.
3. Blocking: prepare 5 % milk or 5% BSA solution in 1 TBST (300 ml for 2 blots); incubate the membrane in 1 methanol for 30 sec, then wash it in 1 TBST; incubate the membrane in the solution for 60 min and wash it twice in 1 TBST.
4. Incubation: Incubate blot with primary antibody at an appropriate concentration (e.g. 1:1000) according to the standard protocol overnight in the cold room (4C) on a nutator (in a seal-a-meal bag); Quickly wash blot twice in 1 TBST, then take 1 15 min and 3 5 min washes in 1 TBST (in a small Tupperware on a shaker); Incubate blot with secondary antibody (rabbit, or mouse, or goat) at an appropriate (e.g. 1:3000) in incubation buffer for 1 hour at room temperature on a nutator (in a seal-a-meal bag); Quickly wash blot twice in 1 TBST, then wash for 2 10 min in 1 TBST.
5. Develop: Place blot on a piece of plastic wrap on your bench; Add ECL on top of the blot for 1 min (use equal parts of solution 1 and solution 2); Shake off excess solution from the blot and wrap securely in a new piece of plastic wrap; Expose blot for 1-5 min.
2.2.10 Crystal Violet Assay
1. Remove the medium by pipetting or dump
2. Be moved the cells with 50 l of 0.5% crystal violet solution per well, he
3. Incubate the cells for 20 min at room temperature on a shaker
4. Wash the plate (s) at least 5-6 times with 200 l water (colored is not knocked out)
5. Tap the remaining water from the wells and dry the cells at room temperature
6. Put 200 l methanol per well and incubate 20 minutes on the shaker at room
7. Exhibition at 590 nm
2.2.11 SiRNA Transfection
1. Prepare a 5 M siRNA solution in RNAse-free water.
2. In separate tubes, dilute the appropriate volume of 5 M siRNA (Tube 1) and the desired DharmaFECT1 transfection reagent (Tube 2) with serum-free medium.
a. Tube 1 - Add 44 L of 5 M siRNA to 66 L RNase-free water.
Add 110 L OptiMEM.
The total volume is 220 L.
b. Tube 2 - Add3.85 L of DharmaFECT 1 to 216.15 OptiMEM.
The total volume is 220 L.
3. Mix the contents of each tube gently by pipetting carefully up and down and incubate for 5 minutes at room temperature.
4. Add contents of Tube 1 to Tube 2.
5. Mix by pipetting carefully up and down and incubate for 20 minutes at room temperature.
6. Remove culture medium from the wells of the 6-well plate and add 400 L of the appropriate transfection mix to each well.
7. Incubate cells at 37oC in 5% CO2 for 72 hrs.
2.2.12 Invasion Assay
1. Coating of the ThinCert? cell culture inserts
a. Using sterile forceps remove 24 well ThinCert? cell culture inserts with 8 m pores and translucent PET membranes from their packaging and pit them into wells of a CELLSTAR? 24 well cell culture plate.
b. Dilute 10 mg Collagen 1 (Roche Diagnostics, GmbH, Mannheim, Germany) with 25 ml steroid cold water (0.2% acetate) to yield a protein concentration of 0.4 mg/ml.
c. Add 50l of diluted Collagen 1 to the center of each cell well inserts. Gently spread the Matrigel across the entire surface of the membrane.
d. Place coated inserts under laminar air flow to allow the Matrigel to solidify for at least 1.5 hours.
e. Non-coated ThinCert? cell culture inserts were used for Migration assay.
2. Preparation of the cells
a. Treat and grow cells according to standard cell culture procedures. The PDAC cell lines were cultured in RPMI medium with 10 % FCS. The night before the migration/invasion experiment the cells were deprived in appropriate free reduced culture medium (0.5% FCS).
b. Cells were harvested and resuspended in 0.5% FCS prior to adjustment to a final concentration of 1 105 per ml.
3. Sowing of the cells and incubation
a. Plate 5 104 cells in 500 l of 0.5% FCS into upper chamber.
b. Add 750 l of 10 % FCS into lower chamber.
c. Culture for approximately 20 hours.
d. Incubate the plate with inserts for 24 hours in a cell culture incubator at 37C and 5 % CO2.
4. Labelling, detachment and quantification of the cells
a. After culture, aspirate medium from upper and lower chambers and wash by 1 PBS once.
b. Gently remove cells that did not migrate through the pores and therefore remained on the upper chamber with a cotton swab (Invaded Cells group). Cleanse the filter as completely as possible and be careful not to press too firmly, the membrane may be popped out. The untreated chambers were considered as the Total Cells group.
c. Add 0.2 ml and 0.4 ml of 1% Crystal Violet in 2% ethanol into upper and lower chamber, respectively, incubate for 30 min.
d. Aspirate Crystal Violet and wash each well 3 times with dH2O (0.5 ml and 1.0 ml for upper and lower chamber, respectively) for 15 minutes each.
e. Add 0.2 ml and 0.3 ml of 10% acetic acid into upper and lower chamber, respectively, incubate for 1.5 hours.
f. Mix the solution in the upper and lower chamber, discard Transwell chamber, and transfer 100 l of the solution from each well of the 24-well plate to a well of a flat-bottom black 96-well plate 2.
g. Finally, quantify the black 96 well plate with a photometer (Sunrise) at an excitation wavelength of 550 nm and an emission wavelength of 700 nm.
5. Calculation of the results
To determine the invasive properties of the studied cell lines the invasion rate (%) was calculated according to the following formula: Invasion Rate (%) = 100 relative fluorescence units obtained from Invaded Cells group / relative fluorescence units obtained from Total Cells group.
2.2.13 Dual Luciferase Activity Assay
1. Reagent Preparation
a. 1 passive lysis buffer (PLB): Add 1 volume of LBP to 4 volumes of distilled water.
b. Luciferase assay reagent (LAR) 2: Resuspend the lyophilized Luciferase assay substrate in Luciferase assay buffer 2.
c. Stop & Glo? Reagent: Add 2.1ml of 50 Stop & Glo? Reagent to 105ml of Stop & Glo? Buffer in the amber Stop & Glo? Reagent bottle provided. Vortex 10 seconds.
2. cell lysis
a. Remove growth media from cultured cells.
b. Rinse cultured cells in 1 PBS. Remove all rinse solution.
c. Dispense the recommended volume of 1PLB into each culture vessel.
3. Dual-Luciferase? Assay step
a. Set injectors 1 and 2 to dispense 100l of LAR 2 and Stop & Glo? Reagent, respectively.
b. For measurements, use a 1- to 2-second delay and a 5- to 10-second read time.
c. Plate with 20l of PLB Lysate/well.
d. Dispense 100l of LAR 2.
e. Measure firefly luciferase activity.
f. Dispense 100l of Stop & Glo? Reagent.
g. Measure Renilla luciferase activity.
h. Firefly luciferase activity was normalized to Renilla luciferase activity.