Introduction

The CloneMiner™ cDNA Library Construction Kit is designed to construct high-quality cDNA libraries without the use of traditional restriction enzyme cloning methods. This novel technology combines the performance of SuperScript™ II Reverse Transcriptase with the Gateway® Technology.
Single-stranded mRNA is converted into double stranded cDNA containing attB sequences on each end. Through site-specific recombination, attB-flanked cDNA is cloned directly into an attP-containing donor vector without the use of restriction digestion or ligation.

The resulting Gateway® entry cDNA library can be screened with a probe to identify a specific entry clone. This clone can be transferred into the Gateway® destination vector of choice for gene expression and functional analysis. Alternatively, the entire entry cDNA library can be shuttled into a Gateway®destination vector to generate an expression library.
 
Features of the CloneMiner™ cDNA Library Construction Kit include:

  • SuperScript™ II reverse transcriptase for efficient conversion of mRNA into cDNA
  • Biotin-attB2-Oligo(dT) Primer for poly(A) mRNA binding and incorporation of the attB2 sequence to the 3' end of cDNA
  • attB1 Adapter for ligation of the attB1 sequence to the 5' end of double-stranded cDNA
  • attP-containing vector (pDONR™222) for recombination with attB-flanked cDNA to produce an entry library through the Gateway®BP recombination reaction


 
Advantages of the CloneMiner™ cDNA Library Construction Kit

Using CloneMiner™ cDNA Library Construction Kit offers the following advantages:

  • Produces high yields of quality, double-stranded cDNA
  • Eliminates use of restriction enzyme digestion and ligation allowing cloning of undigested cDNA
  • Highly efficient recombinational cloning of cDNA into a donor vector results in a higher number of primary clones compared to standard cDNA library construction methods (Ohara and Temple, 2001)
  • Reduces number of chimeric clones and reduces size bias compared to standard cDNA library construction methods (Ohara and Temple, 2001)
  • Enables highly efficient transfer of your cDNA library into multiple destination vectors for protein expression and functional analysis


Experimental Summary

The following diagram summarizes the cDNA synthesis process of the CloneMiner™ cDNA Library Construction Kit.
 




The Gateway®Technology

Gateway®is a universal cloning technology based on the site-specific recombination properties of bacteriophage lambda (Landy, 1989). The Gateway® Technology provides a rapid and highly efficient way to move DNA sequences into multiple vector systems for functional analysis and protein expression. For more information on the Gateway® Technology, see next section.



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Choosing a Library Construction Method

Introduction

There are several ways to construct your cDNA library using the CloneMiner™ cDNA Library Construction Kit. You will need to decide between:

  • Radiolabeling or not radiolabeling your cDNA
  • Size fractionating your cDNA by column chromatography or by gel electrophoresis


We recommend radiolabeling your cDNA and size fractionating your cDNA by column chromatography. This section provides information to help you choose the library construction method that best suits your needs.
 
Radiolabeling vs. Non-Radiolabeling

The table below outlines the advantages and disadvantages of the radiolabeling and non-radiolabeling methods. Use this information to choose one method to construct your cDNA library.

 
Radiolabeling Method
Non-Radiolabeling Method
Analyzing First Strand Synthesis
Direct measure of cDNA yield and overall quality of the first strand
No knowledge of cDNA yield or quality until the library is constructed
Determining cDNA Yields for Cloning
Reliable quantitative method using scintillation counter
Qualitative, subjective method using agarose plate spotting assay
Sensitivity of cDNA Detection
Very sensitive to a wide range of cDNA amounts using scintillation counter
Sensitive in detecting 1-10 ng of cDNA per spot (see Performing the Plate Spotting Assay).
Limited resolution for cDNA yields greater than 10 ng per spot (see Performing the Plate Spotting Assay).
Experimental Time
Time consuming filter washes, counting samples, performing calculations
DNA standards and plates for the plate spotting assay can be prepared in advance for several experiments, limited calculations
Preparation
Requires extensive preparation of reagents, equipment, and work area
Requires minimal preparation of DNA standards and agarose plates for the plate spotting assay
Lab Environment
Need to work in designated areas, dispose of radioactive waste, monitor work area, follow radioactive safety regulations
Regular lab environment with no radioactive hazards or radioactive safety regulations



Be sure to read the section entitled Advance Preparation, to prepare any necessary reagents required for your method of choice. If you will be using the radiolabeling method, also read the section entitled Working with Radioactive Materials. If you will be using the non-radiolabeling method, we recommend that you read the section entitled Performing the Plate Spotting Assay, before beginning.
 
 
Choosing a Size Fractionation Method

Size fractionation generates cDNA that is free of adapters and other low molecular weight DNA. Although we recommend size fractionating your cDNA by column chromatography, you may also size fractionate your cDNA by gel electrophoresis. Either method can be used with radiolabeled or non-radiolabeled cDNA. Refer to the guidelines outlined below and choose the method that best suits your needs.
 
Column Chromatography

Column chromatography is commonly used to size fractionate cDNA. Use the column chromatography method to generate a cDNA library with an average cDNA insert size of approximately 1.5 kb (if you start with high-quality mRNA).
Protocols to size fractionate radiolabeled or non-radiolabeled cDNA by column chromatography are provided in this protocol.
 
Gel Electrophoresis

Use the gel electrophoresis method to generate a cDNA library with a larger average insert size (>2.0 kb) or to select cDNA of a particular size. Protocols to size fractionate radiolabeled or non-radiolabeled cDNA by gel electrophoresis are provided in the CloneMiner™ cDNA Construction Kit Web Appendix. Because you will need to have additional reagents on hand, we recommend reading the Web Appendix before beginning. This manual is available from our Web site (www.invitrogen.com) or by contacting Technical Service.
 

The CloneMiner™ cDNA Library Construction Kit is designed to help you construct a cDNA library without the use of traditional restriction enzyme cloning methods. Use of this kit is geared towards those users who have some familiarity with cDNA library construction. We highly recommend that users possess a working knowledge of mRNA isolation and library construction techniques before using this kit.
 
For more information about these topics, refer to the following published reviews:

  • cDNA library construction using restriction enzyme cloning: see Gubler and Hoffman, 1983 (Gubler and Hoffman, 1983) and Okayama and Berg, 1982 (Okayama and Berg, 1982)
  • cDNA library construction using the l-att recombination system: see Ohara and Temple, 2001 and Ohara et. al., 2002 (Ohara and Temple, 2001) (Ohara, 2002)
  • mRNA handling techniques: see Chomczynski and Sacchi, 1987 (Chomczynski and Sacchi, 1987)

 

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Working with Radioactive Material

Introduction

Read the following section if you will be constructing your cDNA library using a radiolabeled isotope. This section provides general guidelines and safety tips for working with radioactive material. For more information and specific requirements, contact the safety department of your institution.
 
Use extreme caution when working with radioactive material. Follow all federal and state regulations regarding radiation safety. For general guidelines when working with radioactive material, see below.
 
 
General Guidelines

Follow these general guidelines when working with radioactive material.

  • Do not work with radioactive materials until you have been properly trained.
  • Wear protective clothing, gloves, and eyewear and use a radiation monitor.
  • Use appropriate shielding when performing experiments.
  • Work in areas with equipment and instruments that are designated for radioactive use.
  • Plan ahead to ensure that all the necessary equipment and reagents are available and to minimize exposure to radioactive materials.
  • Monitor work area continuously for radiation contamination.
  • Dispose of radioactive waste properly.
  • After you have completed your experiments, monitor all work areas, equipment, and yourself for radiation contamination.
  • Follow all the radiation safety rules and guidelines mandated by your institution.

Any material in contact with a radioactive isotope must be disposed of properly. This will include any reagents that are discarded during the cDNA library synthesis procedure (e.g. phenol/chloroform extraction, ethanol precipitation, cDNA size fractionation). Contact your safety department for regulations regarding radioactive waste disposal.
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Experimental Timeline

Introduction

The CloneMiner™ cDNA Library Construction Kit is designed to produce an entry library from your starting mRNA within three days. It will take an additional two days to determine the titer and quality of the cDNA library. Note that this protocol is organized according to the recommended timeline below. If you will not be following this timeline, be sure to plan ahead for convenient stopping points (see below for more information).
 
Recommended Timeline






If you are performing the radiolabeling method, we recommend that you follow the timeline outlined above. Radiochemical effects induced by 32P decay in the cDNA can reduce transformation efficiencies over time.
 
Optional Stopping Points

If you cannot follow the recommended timeline, you may stop the procedure during any ethanol precipitation step. These steps occur during second strand synthesis and size fractionation and are noted as optional stopping points. When stopping at these points, always store the cDNA as the uncentrifuged ethanol precipitate at -20°C to maximize cDNA stability.

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Experimental Overview

Introduction

The experimental steps necessary to synthesize attB-flanked cDNA and to generate an entry library are outlined below. Once you have isolated your mRNA, you will need a minimum of 3 days to construct a cDNA library.

       Day
         Step
Action
1
1
Synthesize the first strand of cDNA from your isolated mRNA using the Biotin-attB2-Oligo(dT) Primer and SuperScript II RT.
 
2
Synthesize the second strand of cDNA using the first strand cDNA as a template.
 
3
Analyze the first strand reaction for cDNA yield and percent incorporation of [a-32P]dCTP.
 
4
Ligate the attB1 adapter to the 5' end of your cDNA.
2
1
Size fractionate the cDNA by column chromatography to remove excess primers, adapters, and small cDNA.
 
2
Perform the BP recombination reaction between the attB-flanked cDNA and pDONR222.
3
1
Transform the BP reactions into ElectroMAX DH10B T1 Phage Resistant cells. Add freezing media to transformed cells to get final cDNA library.
 
2
Perform the plating assay to determine the cDNA library titer.
4-5
1
Calculate the cDNA library titer using the results from the plating assay.
 
2
Inoculate 24 positive transformants from the plating assay. Determine average insert size and percent recombinants by restriction analysis.
 
3
Sequence entry clones to verify presence of cDNA insert, if desired.

 

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Isolating mRNA

Introduction

You will need to isolate high-quality mRNA using a method of choice prior to using this kit. Follow the guidelines provided below to avoid RNase contamination.
 
Aerosol-resistant pipette tips are recommended for all procedures. See below for general recommendations for handling mRNA.
 
General Handling of mRNA

When working with mRNA:

  • Use disposable, individually wrapped, sterile plasticware
  • Use only sterile, RNase-free pipette tips and RNase-free microcentrifuge tubes
  • Wear latex gloves while handling all reagents and mRNA samples to prevent RNase contamination from the surface of the skin
  • Always use proper microbiological aseptic technique when working with mRNA

You may use RNase Away™ Reagent, a non-toxic solution available from Invitrogen, to remove RNase contamination from surfaces. For further information on controlling RNase contamination, see Current Protocols in Molecular Biology (Ausubel et al., 1994) or Molecular Cloning: A Laboratory Manual (Sambrook et al., 1989).
 
mRNA Isolation

mRNA can be isolated from tissue, cells, or total RNA using the method of choice. We recommend isolating mRNA using the Micro-FastTrack™ 2.0 or FastTrack® 2.0 mRNA Isolation Kits available from Invitrogen. Generally, 1 to 5 µg of mRNA will be sufficient to construct a cDNA library containing 106 to 107 primary clones in E. coli. Resuspend isolated mRNA in DEPC-treated water and check the quality of your preparation. Store your mRNA preparation at -80°C. We recommend aliquoting your mRNA into multiple tubes to reduce the number of freeze/thaw cycles.
 
It is very important to use the highest quality mRNA possible to ensure success. Check the integrity and purity of your mRNA before starting.
 
 
Checking the Total RNA Quality

To check total RNA integrity, analyze 1 µg of your RNA by agarose/ethidium bromide gel electrophoresis. You should see the following on a denaturing agarose gel:

  • 28S rRNA band (4.5 kb) and 18S rRNA band (1.9 kb) for mammalian species
  • 28S band should be twice the intensity of the 18S band

 
Checking the mRNA Quality

mRNA will appear as a smear from 0.5 to 12 kb. rRNA bands may still be faintly visible. If you do not detect a smear or if the smear is running significantly smaller than 12 kb, you will need to repeat the RNA isolation. Be sure to follow the recommendations listed on the previous page to prevent RNase contamination.
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Advance Preparation

Introduction

Some of the reagents and materials required to use the CloneMiner™ cDNA Library Construction Kit are not supplied with the kit and may not be common lab stock. Refer to the lists below to help you prepare or acquire these materials in advance.
 
Refer to the section entitled Before Starting at the beginning of each procedure for a complete list of required reagents.
 
 
Materials Required for the Radiolabeling Method

You should have the following materials on hand before performing the radiolabeling method:

  • [a-32 P]dCTP, 10 µCi/µl (Amersham Biosciences, Catalog no. PB.10205)
  • Glass fiber filters GF/C, 21 mm circles (Whatman, Catalog no. 1822 021)
  • Solvent-resistant marker (Fisher Scientific, Catalog no. 14-905-30)
  • 10% trichloroacetic acid + 1% sodium pyrophosphate (See Recipes)
  • 5% trichloroacetic acid (See Recipes)

 
Materials Required for the Non-Radiolabeling Method

You should have the following on hand before performing the non-radiolabeling method.

  • SYBR® Gold Nucleic Acid Gel Stain, recommended (Molecular Probes, Catalog no. S-11494). Other stains are suitable.

Number of Reactions

This kit provides enough reagents to construct five cDNA libraries. While some reagents are supplied in excess, you may need additional reagents and materials if you wish to perform more than 5 reactions. You may also need additional electrocompetent E. coli cells if you will be performing control reactions (2.3 kb RNA control, pEXP7-tet control, BP negative control, and pUC 19 transformation control) each time you construct a cDNA library. 
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Day 1: Synthesizing cDNA with Flanking attB Sites






Synthesizing the First Strand

Introduction

This section provides detailed guidelines for synthesizing the first strand of cDNA from your isolated mRNA. The reaction conditions for first strand synthesis catalyzed by SuperScript™ II RT have been optimized for yield and size of the cDNAs. To ensure that you obtain the best possible results, we suggest you read this section and the sections entitled Synthesizing the Second Strand and Ligating the attB1 Adapter before beginning.
 

cDNA synthesis is a multi-step procedure requiring many specially prepared reagents which are crucial to the success of the process. Quality reagents necessary for converting your mRNA sample into double-stranded cDNA are provided with this kit. To obtain the best results, do not substitute any of your own reagents for the reagents supplied with the kit.

 
 
Starting mRNA

To successfully construct a cDNA library, it is crucial to start with high-quality mRNA. For guidelines on isolating mRNA. The amount of mRNA needed to prepare a library depends on the efficiency of each step. Generally, 1 to 5 µg of mRNA will be sufficient to construct a cDNA library containing 106 to 107 primary clones in E. coli.
 
2.3 kb RNA Control

We recommend that you include the 2.3 kb RNA control in your experiments to help you evaluate your results. The 2.3 kb RNA control is an in vitro transcript containing the tetracycline resistance gene and its promoter (Tcr).
 
Guidelines

Consider the following points before performing the priming and first strand reactions:

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  • We recommend using no more than 5 µg of starting mRNA for the first strand synthesis reaction
  • Both the amount of DEPC-treated water used to dilute your mRNA and the total volume of your reactions will depend on the concentration of your starting mRNA
  • We recommend using a thermocycler rather than a water bath both for ease and for accurate temperatures and incubation times
  • Tubes should remain in the thermocycler or water bath when adding SuperScript™ II RT to minimize temperature fluctuations (see Hot Start Reverse Transcription, below)


 
Hot Start Reverse Transcription

Components of the first strand reaction are pre-incubated at 45°C before the addition of SuperScript™ II RT. Incubation at this temperature inhibits nonspecific binding of primer to template and reduces internal cDNA synthesis and extension by SuperScript™ II RT. For this reason, it is important to keep all reactions as close to 45°C as possible when adding SuperScript™ II RT.
 
If you are constructing multiple libraries, we recommend making a cocktail of reagents to add to each tube rather than adding reagents individually. This will reduce the time required for the step and will also reduce the chance of error.
 
 
Preparing [a-32 P]dCTP

If you will be labeling your first strand with [a-32 P]dCTP (10 µCi/µl), dilute an aliquot with DEPC-treated water to a final concentration of 1 µCi/µl. Use once and properly discard any unused portion as radioactive waste.
 
Using the Non-Radiolabeling Method

If you prefer to construct a non-radiolabeled cDNA library, perform the following protocols substituting DEPC-treated water for [a-32 P]dCTP.
 
Before Starting

You should have the following materials on hand before beginning. Keep all reagents on ice until needed.
Supplied with kit:

  •  2.3 kb RNA control (0.5 µg/µl) (optional)
  •  DEPC-treated water
  •  Biotin-attB2-Oligo(dT) Primer (30 pmol/µl)
  • 10 mM (each) dNTPs
  •  5X First Strand Buffer
  •  0.1 M DTT
  •  SuperScript™ II RT (200 U/µl)


Supplied by user:

  •  High-quality mRNA (up to 5 µg)
  •  Thermocycler (recommended) or water bath, heated to 65°C
  •  Ice bucket
  •  [a-32 P]dCTP, diluted to 1 µCi/µl (radiolabeling method only)
  •  Thermocycler (recommended) or water bath, heated to 45°C
  •  20 mM EDTA, pH 8.0 (radiolabeling method only)


 
Diluting Your Starting mRNA
In a PCR tube or 1.5 ml tube, dilute your starting mRNA with DEPC-treated water according to the table below. The total volume for your mRNA + DEPC-treated water will vary depending on the amount of starting mRNA.
If you will be using the 2.3 kb RNA control supplied with the kit, add 5 µl of DEPC-treated water to 4 µl of the control mRNA for a total volume of 9 µl and a final mRNA amount of 2 µg.

 

 
µg of starting mRNA
 
Reagent
1
2
3
4
5
Control
mRNA + DEPC-treated water
    10 µl
     9 µl
     8 µl
     7 µl
     6 µl
             9 µl
(4 µl of mRNA + 5 µl of water)



Priming Reaction

  1. To your diluted mRNA (mRNA + DEPC-treated water), add the Biotin-attB2-Oligo(dT) Primer and 10 mM dNTPs according to the following table.
     
    µg of starting mRNA
     
    Reagent
    1
    2
    3
     
    4
    5
    Control
    mRNA + DEPC-treated water
         10 µl
           9 µl
           8 µl
    7 µl
           6 µl
                 9 µl
    Biotin-attB2-Oligo(dT) Primer (30 pmol/µl)
           1 µl
           1 µl
           1 µl
    1 µl
           1 µl
                 1 µl
    10 mM (each) dNTPs             
           1 µl
           1 µl
           1 µl
    1 µl
           1 µl
                 1 µl
    Total Volume
         12 µl
         11 µl
         10 µl
    9 µl
           8 µl
               11 µl

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  2. Mix the contents gently by pipetting and centrifuge for 2 seconds to collect the sample.
  3. Incubate the mixture at 65°C for 5 minutes and cool to 45°C for 2 minutes. During these incubation steps, perform step 1 of the First Strand Reaction, below.



First Strand Reaction

  1. Add the following reagents to a fresh tube. Note: If you will be using the non-radiolabeling method, substitute DEPC-treated water for [a-32 P]dCTP.
         
    5X First Strand Buffer                    4 µl
    0.1 M DTT                                       2 µl
    [a-32 P]dCTP (1 µCi/µl)                    1 µl
          
      
  2. Mix the contents gently by pipetting and centrifuge for 2 seconds to collect the sample.
  3. After the priming reaction has cooled to 45°C for 2 minutes (step 3, above), add the mixture from step 1 to the priming reaction tube. Be careful to not introduce bubbles into your sample. The total volume in the tube should now correspond to the following table:


     
    µg of starting mRNA
     
     
    1
    2
    3
    4
    5
    Control
    Total Volume
          19 µl
          18 µl
          17 µl
          16 µl
          15 µl
          18 µl

      
  4. Incubate the tube at 45°C for 2 minutes.
  5. With the tube remaining in the thermocycler or water bath, carefully add SuperScript II RT according to the following table. Note that this step may be difficult.
     
    µg of starting mRNA
     
     
    1
    2
    3
    4
    5
    Control
    SuperScript II RT (200 U/µl)
          1 µl
          2 µl
          3 µl
          4 µl
          5 µl
          2 µ

    The total volume should now be 20 µl regardless of the amount of starting mRNA.

      
  6. With the tube remaining in the thermocycler or water bath, mix the contents gently by pipetting. Be careful to not introduce bubbles. Incubate at 45°C for 60 minutes.
  7. If you are constructing a radiolabeled cDNA library, proceed to First Strand Reaction Sample, below. If you are constructing a non-radiolabeled cDNA library, proceed to Synthesizing the Second Strand.



 
First Strand Reaction Sample

Follow the steps below to generate a sample for first strand analysis. We recommend analyzing the sample during an incubation step in the second strand reaction.

  1. After the first strand reaction has incubated at 45°C for 60 minutes (step 6, above), mix the contents gently by tapping and centrifuge for 2 seconds to collect the sample.
  2. Add 1 µl of the first strand reaction to a separate tube containing 24 µl of 20 mM EDTA, pH 8.0. Mix gently by pipetting and place on ice until you are ready to analyze the first strand reaction (see Analyzing the First Strand Reaction).
  3. Take the remaining 19 µl first strand reaction and proceed immediately to Synthesizing the Second Strand.


 
Synthesizing the Second Strand

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Introduction

This section provides guidelines for synthesizing the second strand of cDNA. Perform all steps quickly to prevent the temperature from rising above 16°C.
 
Before Starting

You should have the following materials on hand before beginning. Keep all reagents on ice until needed.
Supplied with kit:

  • DEPC-treated water
  • 5X Second Strand Buffer
  • 10 mM (each) dNTPs
  •  E. coli DNA Ligase (10 U/µl)
  •  E. coli DNA Polymerase I (10 U/µl)
  •  E. coli RNase H (2 U/µl)
  •  T4 DNA Polymerase (5 U/µl)
  •  Glycogen (20 µg/µl)


Supplied by user:

  • Ice bucket
  • Thermocycler (recommended) or water bath at 16°C
  • 0.5 M EDTA, pH 8.0
  • Phenol:chloroform:isoamyl alcohol (25:24:1)
  • 7.5 M NH4OAc (ammonium acetate)
  • 100% ethanol
  • Dry ice or a -80°C fr

 

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Day 2: Size Fractionating cDNA by Column Chromatography and Performing the BP Recombination Reaction





Size Fractionating Radiolabeled cDNA by Column Chromatography
 
Introduction

Column chromatography optimizes size fractionation of the cDNA and makes the cloning of larger inserts more probable. Follow instructions closely using the columns supplied with the kit to produce the highest quality library possible.
 
 
Use extreme caution when working with radioactive material. Follow all federal and state regulations regarding radiation safety.
 

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How the Columns Work

Each column provided with the kit contains 1 ml of Sephacryl® S-500 HR resin. This porous resin traps residual adapters and/or small cDNAs (<500 bp) and prevents them from contaminating the library. Larger molecules bypass the resin and elute quickly while smaller molecules are retained within the resin and elute more slowly. Thus, earlier eluted fractions contain larger cDNA fragments than later fractions.
 
If you are constructing more than one cDNA library, only add one cDNA adapter ligation reaction per column.
 
 
Before Starting

You should have the following materials on hand before beginning:

Supplied with kit:

  • cDNA Size Fractionation Columns
  • Glycogen (20 µg/µl)


Supplied by user:

  • Ice bucket
  • Thermocycler (recommended) or water bath, heated to 70°C
  • TEN buffer (10 mM Tris-HCl, pH 7.5; 0.1 mM EDTA; 25 mM NaCl)
  • Scintillation vials
  • Scintillation counter
  • 100% ethanol
  • 7.5 M NH4 OAc (ammonium acetate)
  • Dry ice or -80°C freezer
  • 70% ethanol
  • TE buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)


 
Stopping the Ligation Reaction

   1. Incubate the tube from step 2, at 70°C for 10 minutes to inactivate the ligase.

   2. Place the tube on ice.

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Setting Up the Column

Keep the following points in mind when setting up a fractionation column:

  • Anchor the column securely in a support stand
  • Place a rack containing 1.5 ml tubes below the column
  • The outlet of the column should be 1 to 2 cm above the 1.5 ml tubes
  • You will need to be able to freely move the rack under the column


 
Washing the Column

cDNA size fractionation columns are packed in 20% ethanol which must be completely removed before adding your cDNA sample. Follow the steps below to remove the ethanol from the columns. The washing steps will take approximately 1 hour.

  1. With the column attached to a support stand, remove the top cap first followed by the bottom cap. Allow the ethanol to drain completely by gravity.
  2. Once the column stops dripping, pipette 0.8 ml of TEN buffer into the column and let it drain completely. Refer to the important note below for column specifications.
  3. Repeat the wash step three more times for a total of four washes and 3.2 ml of TEN buffer. Let the column drain until dry. Proceed to Collecting Fractions, below.


 
If the flow rate is noticeably slower than 30-40 seconds per drop, do not use the column. If the drop size from the column is not approximately 25 to 35 µl, do not use the column. The integrity and resolution of the cDNA may be compromised if the column does not meet these specifications.
 
 
Collecting Fractions

When collecting fractions, we recommend wearing gloves that have been rinsed with ethanol to reduce static.

  1. Label 20 sterile 1.5 ml tubes from 1 to 20. Place them in a rack 1 to 2 cm from the bottom of the column with tube 1 under the outlet of the column.
  2. Add 100 µl of TEN buffer to the 50 µl heat-inactivated cDNA adapter ligation reaction from step 1, above. Mix gently by pipetting and centrifuge for 2 seconds to collect the sample.
  3. Add the entire sample to the column and let it drain into the resin bed. Collect the effluent into tube 1.
  4. Move tube 2 under the column outlet and add 100 µl of TEN buffer to the column. Collect the effluent into tube 2. Let the column drain completely. Note:  It is important to make sure all of the effluent has drained from the column before adding each new 100 µl aliquot of TEN buffer.
  5. Beginning with the next 100 µl aliquot of TEN buffer, collect single-drop fractions into individual tubes starting with tube 3. Continue to add 100 µl aliquots of TEN buffer until all 18 tubes (tubes 3-20) contain a single drop.


 
Filling Out the Worksheet: Columns A, B, and C

A worksheet is provided to help you with your data recording (see the Appendix).
 
Using a pipet, measure the volume in each tube. Use a fresh tip for each fraction to avoid cross-contamination. Record this value in column A of the worksheet.

  1. Calculate the cumulative elution volume with the addition of each fraction and record this value in column B.
  2. Identify the first fraction that exceeds a total volume of 600 µl in column B. Do not use this fraction or any subsequent fractions for your cDNA library. Important: These fractions (corresponding to fractions 14 through 20 in the sample worksheet) contain increasing amounts of the attB1 Adapter which will interfere with cloning reactions and will contaminate the library. We recommend discarding these tubes to avoid accidentally using them in the remainder of the protocol.
  3. Place each remaining capped tube directly into a scintillation vial. Do not add scintillation fluid. Obtain Cerenkov counts for each tube and record this value in column C.

 

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Filling Out the Worksheet: Columns D and E

Cerenkov counts will appear above background after approximately 300 µl of total volume (corresponding to fraction 5 in the sample worksheet).

  1. For each fraction in which the Cerenkov counts exceed background, calculate the cDNA yield. Refer to Calculating the Double Strand cDNA Yield, below. Record this value in column D.
  2. Divide each cDNA amount in column D by the fraction volume in column A to determine the cDNA concentration for that fraction. Record this value in column E.


 
Calculating the Double Strand cDNA Yield

Cerenkov counts are approximately 50% of the radioactivity that would be measured in scintillant. Use the specific activity (SA) determined from the first strand reaction sample and the equation below to calculate the yield of double-stranded cDNA. 
 
Amount of ds cDNA (ng)
(Cerenkov cpm) x 2 x (4 pmol dNTP/pmol dCTP) x (1000 ng/ µg ds cDNA)
                SA (cpm/pmol dCTP) x (1515 pmol dNTP/µg ds cDNA)
 
 =  (Cerenkov cpm) x 8
          SA x (1.515)
 
Required cDNA Yield

You will need a final cDNA yield of at least 30 ng to perform the BP recombination reaction. Because you will lose approximately half of your sample during the ethanol precipitation procedure, we recommend that you pool a minimum of 60 ng of cDNA from your fractions. See below for guidelines on selecting and pooling cDNA fractions.
 
Selecting and Pooling cDNA Fractions

The first fraction with detectable cDNA above background level contains the purest and largest cDNAs in the population. Because this fraction often does not contain enough cDNA for cloning, you may need to pool several fractions to reach a minimum of 60 ng of cDNA.

  1. Using the worksheet, determine the cDNA yield in the first fraction containing detectable cDNA above background level.
  2. If the cDNA yield in this fraction is less than 60 ng, add cDNA from subsequent fractions until 60 ng of cDNA is reached.


Note:  The first 60 ng of cDNA from a column will make a library with a larger average insert size compared to a library made from the first 100 ng of cDNA. Use the values in column E to calculate the smallest volume needed from the next fraction to obtain the desired amount of cDNA for cloning.
 
Ethanol Precipitation

  1. To the tube of pooled cDNA, add reagents in the following order:
          Glycogen (20 µg/µl)       1 µl
          7.5 M NH4OAc               0.5 volume (i.e. 0.5 x volume of cDNA)
          100% ethanol                2.5 volume [i.e. 2.5 x (volume of cDNA + NH4OAc)]
       
    Note: 
    You may stop at this point and store the tube at -20°C overnight if necessary.
  2. Place the tube in dry ice or at -80°C for 10 minutes. Centrifuge the sample at +4°C for 25 minutes at 14,000 rpm.
  3. Carefully remove the supernatant while trying not to disturb the cDNA pellet. Add 150 µl of 70% ethanol. Note: Use a Geiger counter to monitor the supernatant for the presence of radioactivity. The majority of the radioactivity should be in the pellet and not in the supernatant.
  4. Centrifuge the sample at +4°C for 2 minutes at 14,000 rpm. Carefully remove the supernatant. Repeat the 70% ethanol wash. Remove as much of the remaining ethanol as possible.
  5. Dry the cDNA pellet in a SpeedVac® for 2-3 minutes or at room temperature for 5-10 minutes.
  6. Resuspend the cDNA pellet in 4 µl of TE buffer by pipetting up and down 30-40 times. Transfer the sample to a fresh tube.Note: Use a Geiger counter to make sure you have resuspended and transferred all of the cDNA pellet. The majority of the radioactivity should be found in the fresh tube and not in the old tube.



 
Calculating the cDNA Yield

  1. Place the capped tube containing the resuspended cDNA from step 6, above, directly into a scintillation vial. Do not add scintillation fluid. Obtain Cerenkov counts.
  2. Determine the cDNA yield using the equation below. Refer to Calculating the Double Strand cDNA Yield.


Amount of ds cDNA (ng) =  (Cerenkov cpm) x 8
                                                     SA x (1.515)
 
What You Should See

You should have a final cDNA yield of approximately 30-40 ng to perform the BP recombination reaction. Using approximately 30-40 ng of cDNA in the BP reaction should produce a library containing 5-10 million clones. If your cDNA yield is less than 30 ng, you may pool additional fractions and ethanol precipitate the cDNA. Resuspend any additional cDNA pellets using the cDNA sample from step 6, above. Once you have the desired amount of cDNA, proceed to Performing the BP Recombination Reaction with Radiolabeled cDNA.
 

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Performing the BP Recombination Reaction with Radiolabeled cDNA
 
Introduction

General guidelines are provided below to perform a BP recombination reaction between your attB-flanked cDNA and pDONR™222 to generate a Gateway® entry library. We recommend that you include a positive control and a negative control (no attB substrate) in your experiment to help you evaluate your results. For a map and a description of the features of pDONR™222, see the Appendix.
 
Resuspending pDONR™222

pDONR™222 is supplied as 6 µg of supercoiled plasmid, lyophilized in TE buffer, pH 8.0. To use, resuspend pDONR™222 plasmid DNA in 24 µl of sterile water to a final concentration of 250 ng/µl.
 
Propagating pDONR™222

If you wish to propagate and maintain pDONR™222, we recommend using Library Efficiency® DB3.1™ Competent Cells (Catalog no. 11782-018) from Invitrogen for transformation. The DB3.1™ E. coli strain is resistant to CcdB effects and can support the propagation of plasmids containing the ccdB gene. To maintain the integrity of the vector, select for transformants in media containing 50 µg/ml kanamycin and 30 µg/ml chloramphenicol.

Note:  DO NOT use general E. coli cloning strains including TOP10 or DH5a™ for propagation and maintenance as these strains are sensitive to CcdB effects. DO NOT use the ElectroMAX™ DH10B™ competent cells provided.
 
Positive Control

pEXP7-tet control DNA is included to use as a positive control for the BP reaction. pEXP7-tet contains an approximately 1.4 kb fragment consisting of the tetracycline resistance gene and its promoter (Tcr) flanked by attB sites. Using the pEXP7-tet fragment in a BP reaction with a donor vector results in entry clones that express the tetracycline resistance gene.
 
Recommended cDNA:pDONR™222 Ratio

For optimal results, we recommend using 30-40 ng of cDNA and 250 ng of pDONR™222 in a 10 µl BP recombination reaction. If the amount of cDNA you will be using is out of this range, make the following changes to the protocol below:

  • Adjust the amount of pDONR™222 such that there is an approximately 1:7 mass ratio of cDNA to pDONR™222
  • If you will be using less than 250 ng of pDONR™222, dilute an aliquot of the vector in order to have a large enough volume to accurately pipette
  • Adjust the amount of TE buffer, pH 8.0 to reach a final volume of 7 µl
  • If you will be using more than 4 µl of cDNA, increase the BP reaction to a final volume of 20 µl

 

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Before Starting

You should have the following materials on hand before beginning. Keep all reagents on ice until needed.
Supplied with kit:

  • pDONR™222, resuspended in sterile water to 250 ng/µl
  • pEXP7-tet control DNA (50 ng/µl)
  • 5X BP Clonase™ Reaction Buffer
  • BP Clonase™ enzyme mix (keep at -80°C until immediately before use)


Supplied by user:

  • attB-flanked cDNA (30-40 ng)
  • TE buffer, pH 8.0 (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)
  • 25°C incubator


 
BP Recombination Reaction

The following protocol uses 30-40 ng of cDNA and 250 ng of pDONR™222 in a 10 µl BP reaction. Use 30 ng of your 2.3 kb RNA control cDNA for the BP reaction. If the attB-flanked cDNA sample is greater than 4 µl, see below for necessary modifications.

  1. Add the following components to a sterile 1.5 ml microcentrifuge tube at room temperature and mix.

     
     
    Component
     
    cDNA Sample
    2.3 kb RNA Control
    BP Negative Control
    BP Positive Control
    attB-flanked cDNA (30-40 ng)
    X µl
    X µl
    --
    --
    pDONR222 (250 ng/µl)
    1 µl
    1 µl
    1 µl
    1 µl
    pEXP7-tet positive control (50 ng/µl)
    --
    --
    --
    0.5 µl
    5X BP Clonase Reaction Buffer
    2 µl
    2 µl
    2 µl
    2 µl
    TE buffer, pH 8.0
    to 7 µl
    to 7 µl
    4 µl
    3.5 µl
  2. Remove the BP Clonase™ enzyme mix from -80°C and thaw on ice (~2 minutes)
  3. Vortex the BP Clonase™ enzyme mix briefly twice (2 seconds each time).
  4. Add 3 ml of BP Clonase™ enzyme mix to each sample. Mix the contents gently by pipetting and centrifuge for 2 se
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Day 3: Transforming Competent Cells





Preparing for Transformation
 
Introduction

Once you have performed the BP recombination reaction, you will inactivate the reaction with proteinase K, ethanol precipitate the cDNA, and transform it into competent E. coli. The ElectroMAX™ DH10B™ T1 Phage Resistant Cells have a high transformation efficiency (>1 x 1010 cfu/µg DNA) making them ideal for generating cDNA libraries. Follow the guidelines below to prepare for the transformation procedure.

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Transformation Control

pUC19 plasmid is included to check the transformation efficiency of ElectroMAX™ DH10B™ T1 Phage Resistant Cells. Transform 10 pg of pUC19 using the protocol.
 
Before Starting

You should have the following materials on hand before beginning:
Supplied with kit:

  • Proteinase K (2 µg/µl)
  • Glycogen (20 µg/µl)
  • pUC19 positive control (10 pg/µl)


Supplied by user:

  • BP recombination reactions (from step 5), previous section
  • Water bath, heated to 37°C
  • Thermocycler or water bath, heated to 75°C
  • Sterile water
  • 7.5 M NH4OAc (ammonium acetate)
  • 100% ethanol
  • Dry ice or a -80°C freezer
  • 70% ethanol
  • 15 ml snap-cap tubes (e.g. Falcon™ tubes)
  • Ice bucket


 
Stopping the BP Recombination Reaction

   1.    To each BP reaction from step 5, add 2 µl of proteinase K to inactivate the BP Clonase™ enzyme mix.

   2.     Incubate the reactions at 37°C for 15 minutes then at 75°C for 10 minutes.

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Ethanol Precipitation

  1. To each tube, add reagents in the following order. Use sterile water. Do not use the DEPC-treated water.     
    Sterile water                            90 µl
    Glycogen (20 µg/µl)                  1 µl
    7.5 M NH4OAc                        50 µl
    100% ethanol                       375 µl
         
    If you performed a 20 µl BP reaction, add 80 µl of sterile water to each tube and add all other reagents as listed above.

    Note:  
    You may stop at this point and store the tube at -20°C overnight if necessary.

      
  2. Place the tube in dry ice or at -80°C for 10 minutes. Centrifuge the sample at +4°C for 25 minutes at 14,000 rpm.
  3. Carefully remove the supernatant while trying not to disturb the cDNA pellet. Add 150 µl of 70% ethanol.
  4. Centrifuge the sample at +4°C for 2 minutes at 14,000 rpm. Carefully remove the supernatant. Repeat the 70% ethanol wash. Remove as much of the remaining ethanol as possible.
  5. Dry the cDNA pellet in a SpeedVac® for 2-3 minutes or at room temperature for 5-10 minutes.
  6. Resuspend the cDNA pellet in 9 µl of TE buffer by pipetting up and down 30-40 times.

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    Preparing the Controls

    You will be dividing your cDNA sample into six aliquots and transforming each aliquot into ElectroMAX™ DH10B™ competent cells. To reduce the amount of work, we recommend that you transform only two aliquots of the 2.3 kb mRNA, BP negative, and BP positive controls and one aliquot of the pUC19 control. Consider the following before preparing the controls:

    • If arcing occurs during electroporation, the sample should be immediately discarded. You will need to repeat the electroporation.
    • You may prepare in advance additional aliquots, tubes, cuvettes, and reagents for any additional electroporations you may have to perform. See recommendations for reducing arcing during electroporation.

     
    Aliquoting Samples

    1.  Label six 1.5 ml tubes for each cDNA library sample. For example, if you are constructing multiple libraries, label tubes for library A: A1, A2, A3, etc.
    2.  Label two 1.5 ml tubes for each of the cDNA library controls (2.3 kb mRNA, BP positive, and BP negative controls). For the pUC19 transformation control, label one 1.5 ml tube.
    3. For each 1.5 ml tube from steps 1 and 2, label a duplicate 15 ml snap-cap tube (e.g. Falcon™ tube).
    4. Aliquot cDNA library samples and controls into the appropriate tubes according to the table below. Place tubes on ice.
       
       
      cDNA Library
      2.3 kb RNA Control
      BP Negative Control
      BP Positive Control
       
      pUC 19 Control
      Number of 1.5 ml Tubes
      6
      2
      2
      2
      1
      Aliquot in Each Tube
      1.5 µl
      1.5 µl
      1.5 µl
      1.5 µl
      1.0 µl

        
    5. Proceed to Transforming ElectroMAX™ DH10B™ T1 Phage Resistant Cells.


     
    Transforming ElectroMAX™ DH10B™ T1 Phage Resistant Cells
     
    Each box of ElectroMAX™ DH10B™ T1 Phage Resistant Cells consists of 5 tubes containing 100 µl of competent cells each. Each tube contains enough competent cells to perform 2 transformations using 50 µl of cells per transformation. Once you have thawed a tube of competent cells, discard any unused cells. Do not re-freeze cells as repeated freezing/thawing of cells may result in loss of transformation efficiency.
     
    Before Starting

    You should have the following materials on hand before beginning:

    Supplied with kit:

    • ElectroMAX™ DH10B™ T1 Phage Resistant Cells (thaw on ice before use)
    • S.O.C. medium (Invitrogen, Catalog no. 15544-034)
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    Supplied by user:

    • Ice bucket
    • 0.1 cm cuvettes (on ice)
    • Electroporator
    • 37°C shaking incubator
    • 15 ml snap-cap tubes (e.g. Falcon™ tubes)
    • Freezing media (60% S.O.C. medium:40% glycerol)

     
    Electroporator Settings

    If you are using the BioRad Gene Pulser® II or BTX® ECM® 630, we recommend the following settings:

    Voltage               2.0 kV
    Resistance        200 W
    Capacity              25 µF

    If you are using another electroporator, you will need to optimize your settings using the pUC19 control DNA provided with the kit. The transformation efficiency of the ElectroMAX™ DH10B™ T1 Phage Resistant Cells should be at least 1 x 1010cfu/µg of pUC19 control DNA.
     
     
    Electroporation

    We recommend that you electroporate your controls first followed by your cDNA samples. This will allow you to troubleshoot any arcing problems before you electroporate your cDNA samples (see recommendation below).

    1. To one tube containing a DNA aliquot, add 50 µl of thawed ElectroMAX™ DH10B™ competent cells. Mix gently by pipetting up and down two times. Be careful to not introduce bubbles into your sample.
    2. Transfer the entire contents of the tube from step 1, above, to a cold 0.1 cm cuvette. Distribute the contents evenly by gently tapping each side of the cuvette. Be careful to not introduce bubbles into your sample.
    3. Electroporate the sample using your optimized setting (see Electroporator Settings). If your sample arcs, discard the sample immediately and repeat the electroporation with another aliquot. You will need to electroporate a minimum of 2 aliquots for the 2.3 kb RNA, BP negative, and BP positive controls and 1 aliquot for the pUC19 control.
    4. Add 1 ml of S.O.C. medium to the cuvette containing electroporated cells. Using a pipette, transfer the entire solution to a labeled 15 ml snap-cap tube.
    5. Repeat steps 1-4 for all sample aliquots.
    6. Shake electroporated cells for at least 1 hour at 37°C at 225-250 rpm to allow expression of the kanamycin resistance marker.
    7. After the one hour incubation at 37°C, pool all cells representing one library into a 15 ml snap-cap tube.
    8. Determine the volume for all cDNA libraries and controls and add an equal volume of sterile freezing media (60% S.O.C. medium:40% glycerol).  Note: Do not add freezing media to the pUC19 control. Mix by vortexing. Keep on ice. This is the final cDNA library.
    9. Remove a 200 µl sample from each library and controls and place in 1.5 ml tubes for titer determination. Keep on ice.
    10. Store cDNA libraries at -80°C. You may divide your library into multiple tubes to reduce the number of freeze/thaw cycles.
    11. Proceed to Performing the Plating Assay.
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    Recommendations

    If you experience arcing during transformation, try one of the following:

    • Make sure the contents are distributed evenly in the cuvette and there are no bubbles.
    • Reduce the voltage normally used to charge your electroporator by 10%.
    • Make sure to ethanol precipitate the BP reaction prior to electroporation to reduce the salt concentration.
    • Dilute the 1.5 µl aliquots with water and divide the sample in two. Electroporate extra samples of competent cells. Make sure that you have enough ElectroMAX™ DH10B™ Cells to perform this troubleshooting step.

     
    Performing the Plating Assay
     
    Before Starting

    You should have the following materials on hand before beginning:

    Supplied by user:

    • cDNA library and control aliquots
    • S.O.C. medium (Invitrogen, Catalog no. 15544-034)
    • LB plates containing 50 mg/ml kanamycin (six for each cDNA library and BP reaction controls, warm at 37°C for 30 minutes)
    • LB plates containing 100 mg/ml ampicillin (two for pUC19 control, warm at 37°C for 30 minutes)

     
    Plating Assay

    1. Serially dilute your sample aliquots with S.O.C. medium according to the table below. For each 1:10 serial dilution, add 100 µl of the sample to 900 µl of S.O.C. medium.
    2. You will be plating your serial dilutions in duplicate. You will need six prewarmed LB plates containing 50 mg/ml kanamycin for each cDNA library, 2.3 kb RNA control, BP negative control, and BP positive control. You will need two prewarmed LB plates containing 100 µg/ml ampicillin for the pUC19 transformation control.
    3. Plate 100 µl of each dilution onto prewarmed LB plates containing the appropriate antibiotic.
    4. Incubate plates overnight at 37°C.
    5. Proceed to Days 4-5: Analyzing the cDNA Library.

     
    cDNA Library
    2.3 kb RNA Control
    BP Negative Control
    BP Positive Control
    pUC 19 Control
    Dilutions
     
              10-2
     
              10-3
     
              10-4
     
           10-2
     
           10-3
     
           10-4
     
           undiluted
     
              10-1
     
              10-2
     
              10-2
     
              10-3
     
              10-4
     
              10-2
     
               --
     
               --
    Amount to Plate of Each Dilution
         2 x 100 µl
    2 x 100 µl
    2 x 100 µl
    2 x 100 µl
    2 x 100 µl
    Total Number of LB + Kan Plates
              6
     
            6
             6       6         --
    Total Number of LB + Amp Plates
              --        --        --        --         2

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Days 4-5: Analyzing the cDNA Library




Determining the cDNA Library Titer
 
Introduction

Guidelines are provided below to determine the titer of your cDNA library.
 
Calculations

   1.    Using the results from the plating assay, and the equation below, calculate the titer for each plate.
          cfu/ml =  colonies on plate x dilution factor
                                volume plated (ml)

   2.    Use the titer for each plate to calculate the average titer for the entire cDNA library.

   3.    Use the average titer and the equation below to determine the total number of colony-forming units. 
          Total CFU (cfu) = average titer (cfu/ml) x total volume of cDNA library (ml)

Note:   If you completed 6 electroporations for your cDNA library, the total volume will be 12 ml. For the controls, you will need to extrapolate the total number of colony-forming units using a total volume of 12 ml.
 
Expected Total CFUs 

In general, a well represented library should contain 5 x 106 to 1 x 107 primary clones. If the number of primary clones is considerably lower for your cDNA library, see Troubleshooting.

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What You Should See

See the table below for expected titers and expected total colony-forming units for the control reactions.
 

 
Control
 
Expected Titer
Expected Volume
 
Expected Total CFUs
2.3 kb RNA control
≥ 1 x 106 cfu/ml
12 ml
≥ 1 x 107 cfu
BP positive control
≥ 1 x 106 cfu/ml
12 ml
≥ 1 x 107 cfu
BP negative control
≤ 0.3% of BP
positive control
12 ml
≤ 0.3% of BP positive control
pUC19 control
≥1 x 1010 cfu/µg DNA
--
--



Qualifying the cDNA Library
 
Introduction

It is important to qualify the cDNA library to determine the success of your cDNA library construction. Determining the average insert size and percentage of recombinants will give you an idea of the representation of your cDNA library.
 
General Molecular Biology Techniques

For help with restriction enzyme analysis, DNA sequencing, and DNA biochemistry, refer to Molecular Cloning: A Laboratory Manual (Sambrook et al., 1989) or Current Protocols in Molecular Biology (Ausubel et al., 1994).
 
Before Starting

You should have the following materials on hand before beginning:

Supplied by user:

  • Restriction enzyme BsrG I and appropriate buffer (New England Biolabs, Catalog no. R0575S)
  • 1 Kb Plus DNA Ladder, recommended (Invitrogen, Catalog no. 12302-011). Other DNA ladders are suitable.
  • Electrophoresis apparatus and reagents


 
Analyzing Transformants by BsrG I Digestion

You will be digesting positive transformants with BsrG I to determine average insert size and percentage of recombinants. BsrG I sites generally occur at a low frequency making it an ideal restriction enzyme to use for insert size analysis. BsrG I cuts within the following sites:

  • attL sites of your entry clone to give you the size of your insert
  • attP sites and ccdB gene in pDONR™222 to distinguish non-recombined pDONR™222


 
Restriction Digest

We recommend that you analyze a minimum of 24 positive clones to accurately determine average insert size and the percentage of recombinants.

  1. Pick 24 colonies from the plating assay and culture overnight in 2 ml LB containing 50 µg/ml of kanamycin.
  2. Isolate plasmid DNA using your method of choice. We recommend using the S.N.A.P.™ MiniPrep Kit (Catalog no. K1900-01) or the PureLink™ 96 Plasmid Purification System (Catalog no. 12263-018) if you will be analyzing multiple libraries at a time.
  3. Digest 300-500 ng of plasmid DNA with BsrG I following the manufacturer’s instructions. Also digest 250 ng of supercoiled pDONR™222 with BsrG I as a control.
  4. Electrophorese samples using a 1% agarose gel. Include a DNA ladder to help estimate the size of your inserts.


 
Expected Digestion Patterns

Use the following guidelines to determine the size of the cDNA inserts.

  • The pDONR™222 control will show a digestion pattern of 3 bands of the following lengths:

     
       2.5 kb
      1.4 kb
      790 bp

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  • Each cDNA entry clone should have a vector backbone band of 2.5 kb and additional insert bands 
  • Make sure to digest enough plasmid DNA to be able to visualize smaller insert bands (<300 bp)
  • Make sure to run the gel long enough to distinguish bands representing insert sizes of approximately 2.5 kb from the 2.5 kb vector backbone band


 
Determining Average Insert Size and % Recombinants

  1. Identify clones containing inserts using the guidelines outlined above.
  2. For clones containing inserts, use the DNA ladder to estimate band sizes. If there are multiple bands for a single cDNA entry clone, add all band sizes to calculate the insert size. Do not include the 2.5 kb vector backbone band in your calculations.
  3. Add together the insert sizes for all clones. Divide this number by the number of clones containing inserts to calculate the average insert size for your cDNA library.
  4. Divide the number of clones containing inserts by the number of clones analyzedto determine the percent recombinants.


 
What You Should See

You should see an average insert size of ≥1.5 kb and at least 95% recombinants for your cDNA library. If the average insert size or percent recombinants of your library clones is significantly lower, the cDNA going into the BP recombination reaction is either of poor quality or is insufficient in quantity.  To troubleshoot any of the cDNA synthesis steps, see Troubleshooting.
 
The Next Step

If you wish to sequence entry clones, proceed to Sequencing Entry Clones.
You may screen your cDNA library to identify a specific entry clone and use this entry clone in an LR recombination reaction with a destination vector to generate an expression clone. Refer to the Gateway® Technology manual to perform an LR recombination reaction using a single entry clone. Alternatively, you may transfer your cDNA library into a destination vector to generate an expression library for functional analysis. For detailed guidelines, refer to Performing the LR Library Transfer Reaction.

 
Sequencing Entry Clones
 
Introduction

You may sequence entry clones generated by BP recombination using any method of choice.
 
Sequencing Primers

To sequence inserts in entry clones derived from BP recombination with pDONR™222, we recommend using the following sequencing primers. Refer to the following table for the location of the primer binding sites.

Forward primer (proximal to attL1)   
M13 Forward (-20): 5' -GTAAAACGACGGCCAG-3'

Reverse primer (proximal to attL2)   
M13 Reverse: 5' -CAGGAAACAGCTATGAC-3'


The M13 Forward (-20) and M13 Reverse Primers (Catalog nos. N520-02 and N530-02, respectively) are available separately from Invitrogen. For other primers, Invitrogen offers a custom primer synthesis service. For more information, visit our Web site (www.invitrogen.com) or contact Technical Service.

Note:   If you experience difficulty using the M13 Reverse Primer to sequence entry clones, we recommend using an alternative reverse primer that hybridizes to the poly A tail of your cDNA insert. Design your reverse primer such that it is 5' -(T 23N-3' where N is A, C, or G.

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General Guidelines

The AT rich attL sites in the entry clones may decrease the efficiency of the sequencing reactions. To optimize your sequencing reactions, we recommend the following:

  • Plasmid DNA sample should be of good quality and purity (OD260/OD280 = 1.7-1.99)
  • During plasmid preparation, elute plasmid using deionized water instead of TE buffer


 
Recombination Region

The recombination region of the entry library resulting from pDONR™222 x attB-flanked cDNA is shown below.
 
Features of the Recombination Region:

  • Restriction sites are labeled to indicate the actual cleavage site.
  • Shaded regions correspond to those DNA sequences transferred from the attB-flanked cDNA into the pDONR™222 vector by recombination. Non-shaded regions are derived from the pDONR™222 vector.
  • Bases 441 and 2686 of the pDONR™222 sequence are marked.



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Appendix: Size Fractionating Non-Radiolabeled cDNA by Column Chromatography

Introduction

Column chromatography optimizes size fractionation of the cDNA and makes the cloning of larger inserts more probable. Follow instructions closely using the columns supplied with the kit to produce the highest quality library possible.
 
Because your cDNA is not labeled with [a-32 P]dCTP, you will need to estimate your cDNA yields using a plate spotting assay. You will be performing this assay throughout the size fractionation procedure. We recommend that you read the section entitled Performing the Plate Spotting Assay, before size fractionating your cDNA.
 
 
How the Columns Work

Each column provided with the kit contains 1 ml of Sephacryl® S-500 HR resin. This porous resin traps residual adapters and/or small cDNAs (<500 bp) and prevents them from contaminating the library. Larger molecules bypass the resin and elute quickly while smaller molecules are retained within the resin and elute more slowly. Thus, earlier eluted fractions contain larger cDNA fragments than later fractions.
 
 
If you are constructing more than one cDNA library, only add one cDNA adapter ligation reaction per column.
 
 
Before Starting

You should have the following materials on hand before beginning:
Supplied with kit:

  • cDNA Size Fractionation Columns
  • Glycogen (20 µg/µl)

Supplied by user:

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  • TEN buffer (10 mM Tris-HCl, pH 7.5; 0.1 mM EDTA; 25 mM NaCl)
  • 100% ethanol
  •  7.5 M NH4OAc (ammonium acetate)
  •  Dry ice or -80°C freezer
  •  70% ethanol
  •  TE buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)

 
Stopping the Ligation Reaction

   1.    Incubate the tube from step 2, at 70°C for 10 minutes to inactivate the ligase.

   2.    Place the tube on ice.

 
Setting Up the Column

Keep the following points in mind when setting up a fractionation column:

  • Anchor the column securely in a support stand
  • Place a rack containing 1.5 ml tubes below the column
  • The outlet of the column should be 1 to 2 cm above the 1.5 ml tubes
  • You will need to be able to freely move the rack under the column
 
Washing the Column

cDNA size fractionation columns are packed in 20% ethanol which must be completely removed before adding your cDNA sample. Follow the steps below to remove the ethanol from the columns. The washing steps will take approximately 1 hour.

  1. With the column attached to a support stand, remove the top cap first followed by the bottom cap. Allow the ethanol to drain completely by gravity.
  2. Once the column stops dripping, pipette 0.8 ml of TEN buffer into the column and let it drain completely. Refer to the important note below for column specifications.
  3. Repeat the wash step three more times for a total of four washes and 3.2 ml of TEN buffer. Let the column drain until dry. Proceed to Collecting Fractions, below.

 
If the flow rate is noticeably slower than 30-40 seconds per drop, do not use the column. If the drop size from the column is not approximately 25 to 35 µl, do not use the column. The integrity and resolution of the cDNA may be compromised if the column does not meet these specifications.

 
 
Collecting Fractions

When collecting fractions, we recommend wearing gloves that have been rinsed with ethanol to reduce static.

  1. Label 20 sterile 1.5 ml tubes from 1 to 20. Place them in a rack 1 to 2 cm from the bottom of the column with tube 1 under the outlet of the column.
  2. Add 100 µl of TEN buffer to the 50 µl heat-inactivated cDNA adapter ligation reaction from step 1. Mix gently by pipetting and centrifuge for 2 seconds to collect the sample.
  3. Add the entire sample to the column and let it drain into the resin bed. Collect the effluent into tube 1.
  4. Move tube 2 under the column outlet and add 100 µl of TEN buffer to the column. Collect the effluent into tube 2. Let the column drain completely. Note: It is important to make sure all of the effluent has drained from the column before adding each new 100 µl aliquot of TEN buffer.
  5. Beginning with the next 100 µl aliquot of TEN buffer, collect single-drop fractions into individual tubes starting with tube 3. Continue to add 100 µl aliquots of TEN buffer until all 18 tubes (tubes 3-20) contain a single drop.

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Filling Out the Worksheet: Columns A and B

A worksheet is provided to help you with your data recording.

  1. Using a pipet, measure the volume in each tube. Use a fresh tip for each fraction to avoid cross-contamination. Record this value in column A of the worksheet.
  2. Calculate the cumulative elution volume with the addition of each fraction and record this value in column B.
  3. Identify the first fraction that exceeds a total volume of 600 µl in column B. Do not use this fraction or any subsequent fractions for your cDNA library.

Important:  These fractions contain increasing amounts of the attB1 Adapter which will interfere with cloning reactions and will contaminate the library. We recommend discarding these tubes to avoid accidentally using them in the remainder of the protocol.
 
Filling Out the Worksheet: Columns C and D

You will be estimating the concentration and yield of your cDNA fractions using the plate spotting assay. Refer to Performing the Plate Spotting Assay for detailed guidelines on preparing the plates and staining the DNA.

  1. Using the DNA Spotting Assay protocol, spot 1 µl of each fraction onto a prewarmed plate.
  2. Record the estimated cDNA concentration of each fraction in column C.
  3. Multiply the cDNA concentration in column C by the fraction volume in column A to determine the amount of cDNA for that fraction. Record this value in column D.

 
Required cDNA Yield

You will need a final cDNA yield of 75 ng to perform the BP recombination reaction. Because you will lose approximately half of your sample during the ethanol precipitation procedure, we recommend that you pool a minimum of 150 ng of cDNA from your fractions. See below for guidelines on selecting and pooling cDNA fractions.
 
If you have previously performed the BP recombination reaction using radiolabeled cDNA, note that the amount of non-radiolabeled cDNA required is greater. This larger amount is due to the difference in scale between quantifying DNA by radioactivity using a scintillation counter and quantifying DNA by the plate spotting assay using the DNA standard. Thus, 30 ng of cDNA as measured by counts is roughly equivalent to 50-100 ng of cDNA as measured by comparison to the DNA standard.
 
Selecting cDNA Fractions

The first fractions containing detectable cDNA by the plate spotting assay contain the purest and largest pieces of cDNA in the population. You will want to use cDNA from these fractions for the BP recombination reaction.
We recommend that you also include the fraction preceding the first fraction with detectable cDNA. This fraction may contain large pieces of cDNA in quantities that are not visible using the plate spotting assay.
 
Pooling cDNA Fractions

You will need to pool fractions together to obtain approximately 150 ng of cDNA. Start with the fraction preceding the first fraction containing detectable cDNA. Add cDNA from subsequent fractions until the desired amount of cDNA is reached.

Note: 
The first 150 ng of cDNA from a column will make a library with a larger average insert size compared to a library made from the first 300 ng of cDNA. Use the values in column C to calculate the smallest volume needed from the next fraction to obtain the desired amount of cDNA for cloning.
 
Ethanol Precipitation
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  1. To the tube of pooled cDNA, add reagents in the following order:

  2.          Glycogen (20 µg/µl)     1 µl
             7.5 M NH4OAc              0.5 volume (i.e. 0.5 x volume of cDNA)
            100% ethanol                2.5 volumes  [i.e. 2.5 x (volume of cDNA +NH4OAc)]
         
    Note:  You may stop at this point and store the tube at -20°C overnight if necessary.

      
  3. Place the tube in dry ice or at -80°C for 10 minutes. Centrifuge the sample at +4°C for 25 minutes at 14,000 rpm.

  4. Carefully remove the supernatant while trying not to disturb the cDNA pellet. Add 150 µl of 70% ethanol.

  5. Centrifuge the sample at +4°C for 2 minutes at 14,000 rpm. Carefully remove the supernatant. Repeat the 70% ethanol wash. Remove as much of the remaining ethanol as possible.

  6. Dry the cDNA pellet in a SpeedVac® for 2-3 minutes or at room temperature for 5-10 minutes.

  7. Resuspend the cDNA pellet in 4.5 µl of TE buffer by pipetting up and down 30-40 times. Transfer the sample to a fresh tube.


 
Preparing Aliquots for the Plate Spotting Assay

   1.   Remove 0.5 µl of your cDNA sample from step 6, above, and add to 4.5 µl of TE buffer to make a 1:10 dilution.

   2.   Remove 2.5 µl of the 1:10 dilution and add to 2.5 µl of TE buffer to make a 1:20 dilution.

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Estimating the cDNA Yield

You will be estimating the concentration and yield of your cDNA sample using the plate spotting assay. Refer to Performing the Plate Spotting Assay, for detailed guidelines on preparing the plates and staining the DNA.

  1. Using the DNA Spotting Assay protocol, spot 1 µl of your 1:10 dilution and 1 µl of your 1:20 dilution onto a prewarmed plate.
  2. Estimate the cDNA concentration of the diluted sample. Multiply this concentration by the dilution factor to get the cDNA concentration of your size fractionated cDNA.
  3. Determine the final cDNA yield by multiplying the cDNA concentration by the total volume in the tube.
  4. You may need to prepare additional dilutions of your samples for the plate spotting assay if your spots appear saturated.


What You Should See

You should have a final cDNA yield of approximately 75-100 ng to perform the BP recombination reaction. Using approximately 75-100 ng of cDNA in the BP reaction should produce a library containing 5-10 million clones. If your cDNA yield is less than 75 ng, you may pool additional fractions and ethanol precipitate the cDNA. Resuspend any additional cDNA pellets using the cDNA sample from step 6, Ethanol Precipitation.
 
Once you have the desired amount of cDNA, proceed to Performing the BP Recombination Reaction with Non-Radiolabeled cDNA. 
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Appendix: Performing the BP Recombination Reaction with Non-Radiolabeled cDNA

Introduction

General guidelines are provided below to perform a BP recombination reaction between your attB-flanked cDNA and pDONR™222 to generate a Gateway® entry library. We recommend that you include a positive control and a negative control (no attB substrate) in your experiment to help you evaluate your results. For a map and a description of the features of pDONR™222.
 
 
Resuspending pDONR™222

pDONR™222 is supplied as 6 µg of supercoiled plasmid, lyophilized in TE buffer, pH 8.0. To use, resuspend pDONR™222 plasmid DNA in 24 µl of sterile water to a final concentration of 250 ng/µl.
 
 
Propagating pDONR™222

If you wish to propagate and maintain pDONR™222, we recommend using Library Efficiency® DB3.1™ Competent Cells (Catalog no. 11782-018) from Invitrogen for transformation. The DB3.1™ E. coli strain is resistant to CcdB effects and can support the propagation of plasmids containing the ccdB gene. To maintain the integrity of the vector, select for transformants in media containing 50 µg/ml kanamycin and 30 µg/ml chloramphenicol.

Note:   DO NOT use general E. coli cloning strains including TOP10 or DH5a™ for propagation and maintenance as these strains are sensitive to CcdB effects. DO NOT use the ElectroMAX™ DH10B™ competent cells provided with this kit.
 

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Positive Control

pEXP7-tet control DNA is included with this kit for use as a positive control for the BP reaction. pEXP7-tet contains an approximately 1.4 kb fragment consisting of the tetracycline resistance gene and its promoter (Tcr) flanked by attB sites. Using the pEXP7-tet fragment in a BP reaction with a donor vector results in entry clones that express the tetracycline resistance gene.
 
 
Before Starting

You should have the following materials on hand before beginning. Keep all reagents on ice until needed.

Supplied with kit:

  • pDONR™222, resuspended in sterile water to 250 ng/µl
  • pEXP7-tet positive control (50 ng/µl)
  • 5X BP Clonase™ Reaction Buffer
  • BP Clonase™ enzyme mix (keep at -80°C until immediately before use)


Supplied by user:

  • attB-flanked cDNA (75-100 ng )
  • TE buffer, pH 8.0 (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)
  • 25°C incubator

 

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BP Recombination Reaction

The following protocol uses 75-100 ng of cDNA and 250 ng of pDONR™222 in a 10 µl BP reaction. If the attB-flanked cDNA sample is greater than 4 µl, see below for necessary modifications.

  1. Add the following components to a sterile 1.5 ml microcentrifuge tube at room temperature and mix.

  2.  
     
    Component
     
    cDNA Sample
    2.3 kb RNA Control
    BP Negative Control
    BP Positive Control
    attB-flanked cDNA (75-100 ng)
    X µl
    X µl
    --
    --
    pDONR222 (250 ng/µl)
    1 µl
    1 µl
    1 µl
    1 µl
    pEXP7-tet positive control (50 ng/µl)
    --
    --
    --
    0.5 µl
    5X BP Clonase Reaction Buffer
    2 µl
    2 µl
    2 µl
    2 µl
    TE buffer, pH 8.0
    to 7 µl
    to 7 µl
    4 µl
    3.5 µl

      
  3. Remove the BP Clonase™ enzyme mix from -80°C and thaw on ice (~2 minutes).

  4. Vortex the BP Clonase™ enzyme mix briefly twice (2 seconds each time).

  5. Add 3 µl of BP Clonase™ enzyme mix to each sample. Mix the contents gently by pipetting and centrifuge for 2 seconds to collect the sample. The total volume in each tube should now be 10 µl. Reminder: Return BP Clonase™ enzyme mix to -80°C immediately after use.

  6. Incubate reactions at 25°C for 16-20 hours. Proceed to Day 3: Transforming Competent Cells.



 
Performing a 20 µl BP Reaction

If you will be using more than 4 µl of cDNA, you may increase the total BP reaction volume to 20 µl. You will need to make the following changes to the above protocol:

  • Add an additional 2 µl of 5X BP Clonase™ Reaction Buffer (4 µl total)
  • Add the appropriate amount of TE buffer to reach a final volume of 14 µl
  • Add 6 µl of BP Clonase™ enzyme mix

 

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Appendix: Performing the Plate Spotting Assay

Introduction

If you are constructing a non-radioactive cDNA library, you will be estimating your cDNA yields using a plate spotting assay. Samples will be spotted on agarose and compared under UV light to spots containing known quantities of DNA. Guidelines are provided below to prepare the plates and to perform the assay.
 
The plate spotting assay is an assay to qualitatively determine the concentration and yield of your cDNA samples. Comparison of samples to the DNA standard is subjective and may vary from person to person. In addition, the plate spotting assay is limited in its range of cDNA detection. While you can detect as little as 1 ng of cDNA using SYBR® Gold Nucleic Acid Gel Stain (see Choosing a Nucleic Acid Stain, below), the assay cannot resolve an unlimited amount of cDNA. Generally, spots containing more than 50 ng of cDNA will appear equally stained under UV light.

 
 
Choosing a Nucleic Acid Stain

DNA may be detected using ethidium bromide or SYBR® Gold Nucleic Acid Gel Stain available from Molecular Probes (Catalog no. S11494). We recommend using SYBR® Gold because it is 10-fold more sensitive than ethidium bromide for detecting DNA in electrophoretic gels.
Ethidium bromide staining requires preparing plates containing agarose plus ethidium bromide. SYBR® Gold staining requires preparing agarose-only plates followed by staining the plate using a SYBR® Gold solution. Guidelines are provided in this section for both stains.
 
 
Using the pEXP7-tet Positive Control

Supercoiled pEXP7-tet DNA is included with the kit as a positive control for the BP recombination reaction. pEXP7-tet can also be used as a DNA standard for the plate spotting assay. The concentration of your cDNA samples can be estimated by comparison under UV light to known quantities of pEXP7-tet DNA.
 

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Number of Plates Needed

You will need two plates per library. One plate will contain each of your fractions and another plate will contain cDNA samples that were pooled and ethanol precipitated.
 
 
Before Starting

You should have the following materials on hand before beginning.

Supplied with kit:

  • pEXP7-tet control DNA (50 ng/µl)


Supplied by user:

  • Polystyrene petri dishes, 100 x 15 mm
  • Ethidium bromide (optional, 10 mg/ml)
  • SYBR® Gold Nucleic Acid Gel Stain (recommended; Molecular Probes Catalog no. S11494)
  • 1% agarose in TAE buffer


 
Preparing Plates

  1. Prepare a 100 ml solution of 1% agarose in 1X TAE buffer. Heat until agarose dissolves and let cool for a few minutes.  If you will be staining your cDNA with ethidium bromide, proceed to step 2. If you will not be using ethidium bromide, skip to step 3.
  2. Add 10 µl of ethidium bromide (10 mg/ml) to the agarose solution for a final concentration of 1 µg/ml. Swirl the solution to mix.
  3. Pour the agarose solution into a petri dish just until the bottom is covered. This will be approximately 15 ml for a 100 x 15 mm plate.
  4. Allow agarose to solidify at room temperature (keep plates in the dark if you are using ethidium bromide). Plates can be stored at +4°C for up to one month. Warm plates to room temperature before use.

 

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Preparing pEXP7-tet Control DNA

Serially dilute pEXP7-tet control DNA in TE buffer to final concentrations of:
           25 ng/µl
           10 ng/µl
             5 ng/µl
             1 ng/µl

DNA standards can be stored at -20°C for up to 1 month.
 
 
Labeling Plates

Using a marker, label plates on the bottom side of the petri dish and indicate where the DNA standards and samples will be spotted (see below).
 
Sample Plates for cDNA Size Fractionation by Column Chromatography





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Guidelines

Consider the following points before performing the DNA plate spotting assay:

  • Warm plates to room temperature before using
  • Do not reuse plates
  • Spot DNA standards and cDNA samples within 10 minutes of each other


 
DNA Spotting Assay

  1. Onto a prewarmed plate, spot 1 µl of each pEXP7-tet control DNA dilution. Avoid touching the agarose with the pipette tip. When the 1 µl aliquot is released, capillary action will pull the small volume from the pipette tip onto the plate surface. Avoid formation of bubbles.
  2. Once the DNA standards are spotted, spot 1 µl of each cDNA sample in a similar fashion.
  3. Allow spots to dry at room temperature for 5-15 minutes.
  4. If you are staining your samples with SYBR® Gold, proceed to Staining Plates with SYBR® Gold, below. If you are staining your samples with ethidium bromide, proceed to the next step.
  5. Remove the lid and visualize the plate under UV light and photograph. Note that the labels and samples will be in the reverse order.
  6. Using the known concentration of the DNA standards, estimate the amount of cDNA in each sample.


 
Staining Plates with SYBR® Gold

  1. Add 5 µl of SYBR® Gold to 50 ml of TAE buffer to make a 1x stain. This solution can be stored in the dark per manufacturer’s instructions.
  2. Remove the plate lid and pour the SYBR® Gold solution over the agarose until the entire plate is covered (approximately 15 ml). Place the plate in a box and wrap in foil to protect the solution from light.
  3. Shake the plate on a lab shaker for 20 minutes.
  4. Discard the stain in the appropriate waste. Air dry the plate.
  5. Remove the lid and visualize the plate under UV light and photograph. Note that the labels and samples will be in the reverse order.
  6. Using the known concentration of the DNA standards, estimate the amount of cDNA in each sample.

 

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Appendix: Performing the LR Library Transfer Reaction

Introduction

Once you have qualified your cDNA library and analyzed entry clones, you can perform the LR recombination reaction to transfer your cDNA library into any Gateway® destination vector of choice. If you will be creating an expression library, you will need to follow the guidelines provided in this section for preparing DNA and for performing the LR recombination reaction.

Alternatively, you may screen your cDNA library to identify a specific entry clone and use this entry clone in an LR recombination reaction with a destination vector to generate an expression clone. Refer to the Gateway® Technology manual to perform a standard LR recombination reaction using a single entry clone.
 
 
Before Starting

You should have the following materials on hand before beginning.
 
Supplied with kit:

  • 30% PEG/Mg solution


Supplied by user:

  • S.N.A.P.™ MidiPrep Kit, recommended (Invitrogen, Catalog no. K1910-01)
  • LB media containing 50 µg/ml kanamycin
  • TE buffer (10 mM Tris-HCl, pH 8.0; 1 mM EDTA)
  • Your cDNA library
  • Destination vector of choice (150 ng/µl)
  • LR Clonase™ enzyme mix (Invitrogen Catalog no. 11791-019)
  • 5X LR Clonase™ Reaction Buffer (supplied with LR Clonase™ enzyme mix)
  • Ice bucket
  • Proteinase K (2 µg/µl) (supplied with LR Clonase™ enzyme mix)
  • Sterile water
  • Glycogen (20 µg/µl)
  • 7.5 M NH4OAc
  • 100% ethanol
  • Dry ice or a -80°C freezer
  • 70% ethanol
  • ElectroMAX™ DH10B™ T1 Phage Resistant Cells or equivalent
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Preparing Double-Stranded DNA

You may prepare plasmid DNA from your cDNA library using your method of choice. We recommend using the S.N.A.P.™ MidiPrep Kit (Catalog no. K1910-01). Consider the following points when preparing your DNA:

  • Inoculate 5 x 106-1 x 107 cfu of your cDNA library into 50 ml of LB containing 50 µg/ml kanamycin
  • Grow the culture to an OD600 of 1.0 (approximately 6 hours)
  • Use TE buffer, pH 8.0 to elute your DNA

 
PEG Precipitation

After you have prepared plasmid DNA from your cDNA library, precipitate the DNA using the 30% PEG/Mg solution provided with the kit.

  1. Precipitate the entire eluate with 0.4 volumes of the 30% PEG/Mg solution. Mix well by pipetting.
  2. Centrifuge at room temperature for 15 minutes at 13,000 rpm. Carefully remove the supernatant.
  3. Dry the pellet at room temperature for 10 minutes. Resuspend the pellet in 50 µl of TE buffer. If you started with less than 5 x 106 clones, resuspend the pellet in less TE buffer.
  4. Determine the DNA yield (see Determining DNA Yield, below).
  5. Dilute the DNA to 25 ng/µl. You will need 50 ng of DNA for one LR recombination reaction. You should have enough DNA to perform several LR recombination reactions, if desired.


 
Determining the DNA Yield

  1. Dilute 5-10 µl of the plasmid DNA sample and read the O.D. using a spectrophotometer at 260 nm.
  2. Determine the concentration using the equation below:   [DNA] = (A260) (0.05 mg/ml) (dilution factor)
  3. Determine the total yield by multiplying the concentration by the volume of DNA.
  4. Dilute the DNA to 25 ng/µl.

  5.  
    LR Library Transfer Reaction

    If you have a positive control plasmid for the LR recombination reaction, we recommend including it in your experiment to help you evaluate your results.

    1. Add the following components to a sterile 1.5 ml microcentrifuge tube at room temperature and mix.

    Component
    Sample
    Negative Control
    Positive Control
    cDNA entry library (25 ng/µl)
    2 µl
    --
    --
    Positive control plasmid (25 ng/µl)
    --
    --
    2 µl
    Destination vector (150 ng/µl)
    3 µl
    3 µl
    3 µl
    5X LR Clonase Reaction Buffer
    4 µl
    4 µl
    4µl
    TE Buffer, pH 8.0
    5 µl
    7 µl
    5 µl
    Total volume
    14 µl
    14 µl
    14 µl

      
  6. Remove the LR Clonase™ enzyme mix from -80°C and thaw on ice (~2 minutes).
  7. Vortex the LR Clonase™ enzyme mix briefly twice (2 seconds each time).
  8. Add 6 µl of LR Clonase™ enzyme mix to each sample. Mix well by vortexing briefly twice (2 seconds each time).  Reminder: Return LR Clonase™ enzyme mix to -80°C immediately after use.
  9. Incubate reactions at 25°C for 16-20 hours.
  10. Add 2 ml of the proteinase K solution to each reaction. Incubate the reactions at 37°C for 15 minutes, then at 75°C for 10 minutes.
  11. Proceed to Ethanol Precipitation, below.

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    Ethanol Precipitation

      
    1. To the LR reaction, add reagents in the following order. Be sure to use sterile water and not DEPC-treated water.


    2.       Sterile water                         80 µl
            Glycogen (20 µg/µl)               1 µl
            7.5 M NH4OAc                      50 µl
            100% ethanol                     375 µl

      Note:  You may stop at this point and store the tube at -20°C overnight if necessary.


    3. Place tube in dry ice or at -80°C for 10 minutes. Centrifuge the sample at +4°C for 25 minutes at 14,000 rpm.
    4. Carefully remove the supernatant trying not to disturb the pellet. Add 150 µl of 70% ethanol.
    5. Centrifuge the sample at +4°C for 2 minutes at 14,000 rpm. Carefully remove the supernatant. Repeat the 70% ethanol wash. Remove as much of the remaining ethanol as possible.
    6. Dry the DNA pellet in a SpeedVac® for 2-3 minutes or at room temperature for 5-10 minutes.
    7. Resuspend the DNA pellet in 9 µl of TE buffer by pipetting up and down 30-40 times.

       
      Transforming Competent E. coli

      You may use any recA, endA E. coli strain including TOP10, DH5a™, DH10B™ or equivalent for transformation. Do not transform the LR reaction mixture into E. coli strains that contain the F' episome (e.g. TOP10F'). These strains contain the ccdA gene and will prevent negative selection with the ccdB gene. We recommend using ElectroMAX™ DH10B™ T1 Phage Resistant Cells for maximum transformation efficiency. If you will be using ElectroMAX™ DH10B™ T1 Phage Resistant cells, follow the guidelines outlined in the section entitled Transforming Competent Cells.
       
       
      Analyzing the Expression Library

      Follow the guidelines outlined in the section entitled Analyzing the cDNA Library, to determine the titer, average insert size, and percent recombinants of your expression library. We recommend that you:

      • Analyze transformants by digesting with BsrG I which cuts within both attB sites of the expression library as well as within the attR sites and ccdB gene for non-recombined destination vectors
      • Digest and electrophorese your destination vector with no insert to determine the background BsrG I digestion pattern for your particular destination vector

       
      What You Should See

      When starting with ≥5 x 106 cfu from your cDNA entry library, you should obtain 5 x 106 – 1 x 107 primary clones from one LR recombination reaction. If the number of primary clones is considerably lower for your expression library, you may perform additional LR recombination reactions using any remaining plasmid DNA from your entry library.
      The average insert size and percentage of recombinants of your expression library should be maintained from your cDNA entry library.
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Troubleshooting

The following table lists some potential problems and possible solutions that may help you troubleshoot various steps during cDNA library construction.

Note that the starting mRNA quality is a key factor that will affect the outcome of your results. 

Problem Cause Solution
Low cDNA yield or low incorporation of [a-32P]dCTP after first strand synthesis (radiolabeling method only)Insufficient starting mRNAQuantitate the mRNA by measuring the A260, if possible. We recommend using 1-5 mg of starting mRNA.

Poorly prepared mRNA or degraded mRNAFollow the recommendations for mRNA isolation and working with mRNA.

Old [a-32P]dCTP or [a-32P]dCTP not addedDo not use [a-32P]dCTP that is more than 2 weeks old. Use fresh [a-32P]dCTP. See guidelines on preparing [a-32P]dCTP.

Essential reagent accidentally not added or not workingPerform the 2.3 kb RNA control reaction to verify that the correct reagents have been added and are working properly.

Inaccurate incubation temperatures or temperature fluctuationsPerform the first strand reaction at 45°C. Keep reactions at 45°C when adding SuperScript™ II RT.

SuperScript™ II RT stored incorrectlyStore SuperScript™ II RT at -20°C in a frost-free freezer.
Low cDNA yield after size fractionation by column chromatographyFaulty columnsCheck each column to verify that it is working properly.

Samples run too quickly over columnsLet columns drain completely before adding additional buffer.
Low cDNA library titer with pUC19 transformation control working properlycDNA of poor qualityMake sure the first strand reaction shows >15% percent incorporation of [a-32P]dCTP (radiolabeling method only).

Insufficient ligation of attB1 AdapterPerform the 2.3 kb RNA control reactions to verify the ligation step worked properly.

Incorrect ratio of cDNA to pDONR™222Refer to the recommended ratio of cDNA to pDONR™222 for the BP reaction.
Low cDNA library titer with pUC19 transformation control working properly, continuedInsufficient amount of cDNA used in the BP recombination reactionUse the minimum amount of cDNA required for the BP recombination reaction. Refer to the radiolabeling method and the non-radiolabeling method.

BP Clonase™ enzyme mix is inactive or suggested amount was not used
  • Perform the pEXP7-tet positive control reactions to verify that BP Clonase™ enzyme mix is active

  • Test another aliquot of the BP Clonase™ enzyme mix

  • Make sure that you store the BP Clonase™ enzyme mix at -80°

  • Do not freeze/thaw the BP Clonase™ enzyme mix more than 10 times

  • Use the recommended amount of BP Clonase™ enzyme mix
Few or no colonies obtained from the pUC19 transformation controlRecombination reactions were not treated with proteinase KTreat reactions with proteinase K before transformation.

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Recipes

10% Trichloroacetic Acid + 1% Sodium Pyrophosphate

100% trichloroacetic acid (TCA) (see below)
Sodium pyrophosphate decahydrate

   1.   Dissolve 10 g of sodium pyrophosphate in 750 ml of deionized water.

   2.   Add 100 ml of 100% trichloroacetic acid (TCA).

   3.   Bring final volume to 1 L with deionized water.

   4.   Store at +4°C for up to 6 months.

 
100% Trichloroacetic Acid

This recipe is designed to hydrate one standard 500 g bottle of TCA crystals. If you wish to hydrate a different size bottle of TCA, adjust the volume sizes accordingly. Use caution when handling TCA. TCA causes severe burns and is harmful if swallowed or inhaled.

   1.   Add 227 ml of deionized water to a 500 g bottle of TCA.

   2.   Cap the bottle tightly and invert the bottle slowly several times to dissolve the TCA thoroughly.

   3.   Add a stir bar to the bottle and stir the solution until homogeneous. No further volume adjustment is required.

   4.   Store at room temperature for up to one year.

 
5% Trichloroacetic Acid

   1.   Add 50 ml of 100% trichloroacetic acid to 950 ml of deionized water.

   2.   Store at room temperature for up to 3 months.

 
Freezing Media

60% S.O.C. medium:40% glycerol

   1.   Combine 60 ml of S.O.C. medium and 40 ml of glycerol and stir until solution is homogeneous.

   2.   Autoclave for 30 minutes on liquid cycle.

   3.   Store at room temperature for up to 1 month.
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Sample cDNA Library

Introduction

In this section, we provide a sample experiment to illustrate the cDNA library construction process. This experiment starts with isolated mRNA and continues through construction and qualification of a radiolabeled cDNA library. All steps were performed according to these protocols.
 
Starting mRNA

3 µg of high-quality HeLa cell mRNA
 
First Strand Analysis

A sample of the first strand reaction was removed and analyzed to determine specific activity, cDNA yield, and percent incorporation of [a-32P]dCTP. The unwashed and washed filters gave the following counts:
 
Specific Activity
 
The specific activity was determined using the counts for the unwashed filter and the equation below:

 
Counts per Minute (cpm)   
Unwashed Filter
45998
Washed Filter
2601



SA (cpm/pmol dCTP) = (cpm unwashed filter/10 µl
                                              200 pmol dCTP/10 µl

                        =       45998 cpm/10 µl          
                               200 pmol dCTP/10 µl
                        =     230 cpm/pmol dCTP

First Strand cDNA Yield

The first strand cDNA yield was determined using the counts for the washed filter, the calculated specific activity, and the equation below:
 
cDNA Yield (µg)
=    (cpm of washed filter) x (25 µl/10 µl) x (20 µl/1 µl) x (4 pmol dNTP/pmol dCTP)
                        SA (cpm/pmol dCTP) (3030 pmol dNTP/ µg cDNA)

=    (cpm of washed filter) x 50 x  (4 pmol dNTP/pmol dCTP)
           SA (cpm/pmol dCTP) (3030 pmol dNTP/ µg cDNA)

=    (cpm of washed filter) x (200)
                  SA x (3030)

=      2601 x 200
         230 x 3030

=      0.746 µg cDNA

Percent Incorporation

The percent incorporation of [a-32P]dCTP was determined using the calculated first strand cDNA yield and the equation below:
 
Percent Incorporation =         cDNA yield (µg) x 100    
                                            starting mRNA amount (µg)

                                               =     0.746 µg cDNA x 100
                                                      3 µg starting mRNA
                                               =     25%

The results of the first strand analysis are summarized below:

Specific Activity                 230 cpm/pmol dCTP
cDNA Yield                        0.746 µg
Percent Incorporation       25%
 
 
Size Fractionation by Column Chromatography

After attB1 adapter ligation, the cDNA was size fractionated using column chromatography. The results are listed in the sample worksheet below. Tube 5 was the first tube to give Cerenkov counts above background. Using the data for tube 5, we demonstrate below how the worksheet was filled out.
 
Tube 5 Example                                                                                                                                                                     

The volume in tube 5 was measured to be 36 µl (column A). Adding this volume to the previous cumulative volume (i.e. 306 µl) gave a total volume of 342 µl (column B). The Cerenkov count was 213 cpm (column C).
The double strand cDNA yield was determined using the count value from column C, the specific activity already calculated in the first strand analysis, and the equation below:

Amount of ds cDNA (ng)  
=     (Cerenkov cpm) x 2 x (4 pmol dNTP/pmol dCTP) x (1,000 ng/ µg ds cDNA)
                         SA (cpm/pmol dCTP) x (1515 pmol dNTP/ µg ds cDNA)
 
=     (Cerenkov cpm) x 8
                SA x (1.515)

=       213 x 8   
      230 x 1.515

= 4.9 ng cDNA (column D)
 
The concentration of cDNA was determined using the calculated cDNA yield and the value in column A.

Concentration of cDNA (ng/µl) =  amount of cDNA (ng)
                                                           fraction volume (µl)
 
                                                        =  Column D
                                                            Column A
 
                                                        =   4.9 ng
                                                             36 µl
 
                                                        =   0.136 ng/µl (column E)

Sample cDNA Library Worksheet

 
 
 
Tube
A
Fraction Volume (µl)
B
Total Volume (µl)
C
Cerenkov Counts (cpm)
D
Amount of cDNA
(ng)
E
Concentration of cDNA (ng/µl)
115115122----
28523614----
33427025----
43630615----
5363422134.90.136
634376113626.10.77
735411262860.31.72
836447411494.52.625
9364834427101.62.82
1033516361483.02.52
1136552294767.71.88
1236588213949.11.36
1336624176140.41.12
1436    
1536    
1635    
1736    
1836    
1936    
2036    


Selecting and Pooling Fractions

Fractions 5, 6, and part of fraction 7 were pooled together for a total of 61.1 ng of cDNA (see table below).

Fraction Pooled Volume (µl)Concentration of cDNA (ng/µl)Amount of cDNA (ng)
5360.1364.9
6340.7726.1
717.51.7230.1
Total Pooled cDNA (ng)61.1


Calculating the cDNA Yield

After ethanol precipitation, the pooled cDNA gave a Cerenkov count of 1538 cpm. cDNA yield was determined using the count value, th

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Sample Size Fractionation with Non-Radiolabeled cDNA

Size Fractionation by Column Chromatography

A sample plate and worksheet is provided below to demonstrate how to estimate the yield of your non-radiolabeled cDNA. Samples were size fractionated by column chromatography and cDNA yields were estimated using the plate spotting assay. Refer to Labeling Plates, in Performing the Plate Spotting Assay to see how plates were labeled. Note that samples are in the reverse order.




Serial dilutions of pEXP7-tet control DNA and column fractions 1-13 were spotted and stained with SYBR® Gold as described.

 
 
Tube
A
Fraction Volume (µl)
B
Total Volume (µl)
C
Concentration of cDNA (ng/µl)
D
Amount of cDNA (ng)
1151151----
285236----
334270----
436306----
5363420.518
6343764136
7354118280
83644710360
936483----
1033516----
1136552----
1236588----
1336624----
1436   
1536   
1635   
1736   
1836   
1936   
2036   



Selecting and Pooling Fractions


Fractions 5, 6, and part of fraction 7 were pooled together for a total of 294 ng of cDNA (see table below).

Fraction Pooled Volume (µl) Concentration of cDNA (ng/µl) Amount of cDNA (ng)
5360.518
6344136
717.58140
Total Pooled cDNA (ng)  294



Estimating the cDNA Yield

After ethanol precipitating the pooled cDNA, cDNA yield was estimated using the plate spotting assay. Refer to Labeling Plates, in Performing the Plate Spotting Assay to see how plates were labeled. Note that samples are in the reverse order.






Serial dilutions of pEXP7-tet control DNA and two dilutions of ethanol-precipitated cDNA were spotted and stained with SYBR® Gold as described.

  1:10 Dilution 1:20 Dilution
cDNA Concentration of Diluted Sample (ng/µl)52.5
Final cDNA concentration (ng/µl)5050
Volume of cDNA (µl)44
Total cDNA Yield (ng) 200 200


BP Recombination Reaction

3 µl of the cDNA sample containing a total of 150 ng of cDNA was used in the BP recombination reaction.

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Experimental Worksheet for the Radiolabeling Method

Introduction

A worksheet is provided to help you with your record keeping and calculations. Before you record any data, we suggest you make several copies of this worksheet for use with additional cDNA synthesis reactions.
 

 
 
 
Tube
A
Fraction Volume (µl)
B
Total Volume (µl)
C
Cerenkov Counts (cpm)
D
Amount of cDNA
(ng)
E
Concentration of cDNA (ng/µl)
1
 
 
 
 
 
2
 
 
 
 
 
3
 
 
 
 
 
4
 
 
 
 
 
5
 
 
 
 
 
6
 
 
 
 
 
7
 
 
 
 
 
8
 
 
 
 
 
9
 
 
 
 
 
10
 
 
 
 
 
11
 
 
 
 
 
12
 
 
 
 
 
13
 
 
 
 
 
14
 
 
 
 
 
15
 
 
 
 
 
16
 
 
 
 
 
17
 
 
 
 
 
18
 
 
 
 
 
19
 
 
 
 
 
20
 
 
 
 
 

 

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Experimental Worksheet for the Non-Radiolabeling Method

Introduction

A worksheet is provided to help you with your record keeping and calculations. Before you record any data, we suggest you make several copies of this worksheet for use with additional cDNA synthesis reactions.

 
 
Tube
A
Fraction Volume (µl)
B
Total Volume (µl)
C
Concentration of cDNA (ng/µl)
D
Amount of cDNA (ng)
1
 
 
 
 
2
 
 
 
 
3
 
 
 
 
4
 
 
 
 
5
 
 
 
 
6
 
 
 
 
7
 
 
 
 
8
 
 
 
 
9
 
 
 
 
10
 
 
 
 
11
 
 
 
 
12
 
 
 
 
13
 
 
 
 
14
 
 
 
 
15
 
 
 
 
16
 
 
 
 
17
 
 
 
 
18
 
 
 
 
19
 
 
 
 
20
 
 
 
 

 

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Vector Maps

Features of the Vector

pDONR™222 (4718 bp) contains the following elements. All features have been functionally tested.

Feature
Benefit
rrnB T1 and T2 transcription terminators
Protects the cloned gene from expression by vector-encoded promoters, thereby reducing possible toxicity (Orosz et al., 1991).
M13 forward (-20) priming site
Allows sequencing in the sense orientation.
attP1 and attP2 sites
Bacteriophage λ-derived DNA recombination sequences that permit recombinational cloning of attB-containing cDNA (Landy, 1989).
BsrG I restriction sites
Allows detection and size determination of cDNA inserts by restriction enzyme analysis.
ccdB gene
Allows negative selection of the plasmid.
Chloramphenicol resistance gene
Allows counterselection of the plasmid.
M13 reverse priming site
Allows sequencing in the anti-sense orientation.
Kanamycin resistance gene
Allows selection of the plasmid in E. coli.
pUC origin
Allows high-copy replication and maintenance of the plasmid in E. coli.

 

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References

  1. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K. (1994). Current Protocols in Molecular Biology (New York: Greene Publishing Associates and Wiley-Interscience).
  2. Bernard, P., and Couturier, M. (1992). Cell Killing by the F Plasmid CcdB Protein Involves Poisoning of DNA-Topoisomerase II Complexes. J. Mol. Biol. 226, 735-745.
  3. Bushman, W., Thompson, J. F., Vargas, L., and Landy, A. (1985). Control of Directionality in Lambda Site Specific Recombination. Science 230, 906-911.
  4. Chomczynski, P., and Sacchi, N. (1987). Single Step Method of RNA Isolation by Acid Guanidinium Thiocyanate-Phenol-Chloroform Extraction. Anal. Biochem. 162, 156-159.
  5. Gubler, U., and Hoffman, B. J. (1983). A Simple and Very Efficient Method for Generating cDNA Libraries. Gene 25, 263-269.
  6. Landy, A. (1989). Dynamic, Structural, and Regulatory Aspects of Lambda Site-specific Recombination. Annu. Rev. Biochem. 58, 913-949.
  7. Ohara, O., Nagase, T., Mitsui, G., Kohga, H., Kikuno, R., Hiraoka, S., Takahashi, Y., Kitajima, S., Saga, Y., and Koseki, H. (2002). Characterization of Size-Fractionated cDNA Libraries Generated by the in vitro Recombination-Assisted Method. DNA Res. 9, 47-57.
  8. Ohara, O., and Temple, G. (2001). Directional cDNA Library Construction Assisted by the in vitro Recombination Reaction. Nucleic Acids Res. 29, e22.
  9. Okayama, H., and Berg, P. (1982). High-Efficiency Cloning of Full-Length cDNA. Mol. Cell. Biol. 2, 161-170.
  10. Orosz, A., Boros, I., and Venetianer, P. (1991). Analysis of the Complex Transcription Termination Region of the Escherichia coli rrnB Gene. Eur. J. Biochem. 201, 653-659.
  11. Ptashne, M. (1992). A Genetic Switch: Phage (Lambda) and Higher Organisms (Cambridge, MA: Cell Press).
  12. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, Second Edition (Plainview, New York: Cold Spring Harbor Laboratory Press).
  13. Weisberg, R. A., and Landy, A. (1983) Site-Specific Recombination in Phage Lambda. In Lambda II, R. A. Weisberg, ed. (Cold Spring Harbor, NY: Cold Spring Harbor Press), pp. 211-250.

 

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MAN0001697       11-6-2009