Southern with DIG Probe

1. BACKGROUND INFORMATION FOR SOUTHERN BLOT ANALYSIS

Southern blotting has been one of the cornerstones of DNA analysis since its first description by E.M. Southern (1975). Southern was the first to show that immobilization of size-fractionated DNA fragments could be carried out in a reliable and efficient manner. The advent of Southern transfer and the associated hybridization techniques made it possible for the first time to obtain information about the physical organization of single and multicopy sequences in complex genomes. The term Southern blotting, now used to describe any type of DNA transfer from gel to membrane.

1.1. Critical Parameters

The amount of DNA that must be loaded depends on the relative abundance of the target sequence to which hybridization will take place. The detection limit of a radioactive probe with a specific activity of 108 to 109 dpm/µg is about 0.5 pg DNA. Thus, for human genomic DNA, 10mg – equivalent to 1.5 pg of a single-copy gene 500 bp in length – is a reasonable minimum quantity to load. Similar sensitivities can be achieved today with nonradioactive probes (see DIG system later).

Optimization of the parameters that influence Southern blotting must be carried out in conjunction with hybridization analysis, as the efficiency of transfer can be assessed only from the appearance of the x-ray film obtained after the membrane is probed. The following paragraphs describe the most important factors that should be considered.

1.2. Choice of membranes

Originally nitrocellulose membranes were used but their fragility prompted to search for alternative types of support matrix, resulting in the introduction of nylon membranes in the early 1980s. The main advantages of nylon membranes is their greater tensile strength and that DNA can be bound covalently by UV cross-linking. Nylon membranes can therefore be reprobed up to about 12 times without becoming broken or losing their bound DNA. In contrast nitrocellulose membranes are fragile (allow only 3 times reprobing) and do not bind DNA covalently: baking to immobilize the DNA leas to a relatively weak hydrophobic attachment through exclusion of water. The disadvantage of nylon membranes is the amount of background signal seen after hybridization. On the other site nylon membranes are able to bind about five times more DNA per cm2 than nitrocellulose. Nylon retains DNA fragments down to 50 nucleotides in length, but nitrocellulose is inefficient with molecules < 500 nucleotides. Hence nitrocellulose is not a good choice for restriction-digested genomic DNA or PCR fragments if the target band for hybridization probing is likely to be < 500 bp.

1.3. Transfer buffer

With nitrocellulose the transfer buffer must provide a high ionic strength to promote binding of the DNA to the membrane. Several formulations exists but 20 x SSC is recommended because it is easy to make up and can be stored for several months at room temperature. Lower SSC concentrations (e.g. 10x) should not be used with nitrocellulose as the lower ionic strength may result in loss of smaller DNA fragments during transfer. Alkaline transfer is not suitable for nitrocellulose as they do not retain DNA at pH 9.0 and fall apart after long exposure to alkali.

Nylon membranes are able to bind DNA under a variety of conditions (acid, neutral, alkaline, high or low ionic strength) but a high-salt buffer such as 20x or 10x SSC or SSPE appears to be beneficial. Positively charged nylon membranes have an additional advantage as they bind DNA covalently after transfer in an alkaline buffer (0.4 M NaOH). The only problem with alkaline blotting is that high backgrounds may result if a chemiluminescent detection system is used.

1.4. Duration of transfer

The most difficult parameter to evaluate is duration of transfer. In a capillary system, rat of transfer depends on the size of the DNA, thickness of the gel, and agarose concentration. Upward capillary transfer is slow, the architecture of the blot crushes the gel and retards diffusion of the DNA. With a high-salt buffer, it takes appr. 18 hrs to obtain acceptable transfer of a 15 kb molecule from a 5 mm thick 0.7% agarose gel; with the same gel 90% of the 1 kb molecules will be transferred in 2 hrs. This problem is partially alleviated by the depurination step, which breaks larger molecules in to fragments 1 to 2 kb in length, thereby reducing the time needed for their transfer. If the gel is thicker than 5 mm or has an agarose concentration > 1 %, it cannot be assumed that the larger fragments will have transferred to a sufficient extent after 12 hr.

Alkaline blots are more rapid, with most of the DNA being transferred during the first 2 hrs. More rapid transfer can be achieved with the downward capillary blot procedure

1.5. Transfer method

Capillary transfer is still the most popular method of Southern blotting. Its advantages are simplicity, reliability and the lack of special equipment requirements. Alternative transfer methods include electroblotting, originally developed for protein transfer. This method does not work with high-salt buffers and so is less suitable for nitrocellulose.

The second alternative to capillary transfer is vacuum or positive pressure blotting. The vacuum/pressure must be controlled carefully to avoid compressing the gel and retarding transfer.

1.6. Immobilization techniques

The aim of immobilization is to attach as much of the transferred DNA as possible to the membrane as tightly as possible. The standard methods for nitrocellulose is baking in a vacuum, for nylon membranes – UV irradiation and for positively charged nylon membranes, immobilization during alkaline transfer or UV irradiation.

Baking and alkaline transfer are straightforward procedures that cannot be improved on if carried out according to the instructions. UV cross-linking is more variable. Calibration of the UV source is essential since to long UV treatment can result in a loss of DNA.

2. BACKGROUND FOR HYBRIDIZATION ANALYSIS

The principle of hybridization analysis is that a single-stranded DNA or RNA molecule of defined sequence (the "probe") can base-pair to a second DNA or RNA molecule that contains a complementary sequence (the "target"). The stability of the hybrid depends on the extent of base pairing that occurs. The analysis is usually carried out with a probe that has been labeled and target DNA that has been immobilized on a membrane support. The two critical parameters are sensitivity (sufficient probe must anneal to the target to produce a detectable signal after exposure) and specificity (the probe must be attached only to the desired target sequence).

2.1. Factors influencing sensitivity

The sensitivity of hybridization analysis is determined by how many labeled probe molecules attach to the target DNA. The greater the number of labeled probe molecules that anneal, the greater is the intensity of the hybridization signal. The specific activity is a measure of the incorporation of labeled nucleotides/probe. Modern labeling procedures as nick translation, random priming, PCR or in vitro RNA synthesis routinely provide probes with high specific activity.

The amount of target DNA is also essential and was already mentioned under chapter 1.1. The kind of labeling is another factor influencing sensitivity. Traditionally 32P or 35S labeled probes were used. Today nonradioactive probes are becoming increasingly popular as they have many advantages (see chapter 3).

2.2. Factors influencing specificity

The hybridization incubation is carried out in a high-salt solution that promotes base-pairing between probe and target sequences. Hybridization is normally carried out below the Tm for the probe/target and the specificity of the experiment is the function of post-hybridization washes. The critical parameters are the ionic strength of the final wash solution and the temperature at which this wash is done.

The highly stringent wash conditions should destabilize all mismatched heteroduplexes, so that hybridization signals are obtained only from sequences that are perfectly homologous to the probe.

2.3. Factors influencing hybrid stability and hybridization rate

A.) Hybrid stability

Ionic strength

Tm increases 16.6 °C for each 10-fold increase in monovalent cations between 0.01 and 0.4 M NaCl.

base composition

AT are less stable than GC base pairs in aqueous solutions containing NaCl

destabilizing agents

Each 1% formamide reduces the Tm by about 0.6 °C for a DNA-DNA hybrid. 6 M urea reduces the Tm by about 30 °C.

mismatched base pairs

Tm (melting temperature) is reduced by 1°C for each 1% of mismatching

B.) Hybridization rate

temperature

Maximum rate occurs at 20-25°C below the Tm for DNA-DNA hybrids, 10-15°C below Tm for DNA-RNA hybrids

Ionic strength

Optimal hybridization rate at 1.5 M Na+

destabilizing agents

50% formamide has not effect, but higher or lower concentrations reduce the hybridization rate

mismatched base pairs

each 10% of mismatching reduces the hybridization rate by a factor of two

duplex length

hybridization rate is directly proportional to duplex length

viscosity

increased viscosity increases the rate of membrane hybridization, 10% dextran sulfate increases rate by factor ten.

probe complexity

repetitive sequences increase the hybridization rate

base composition

little effect

pH

little effect between pH 5.0 and pH 9.0.

3. GENERAL CONSIDERATION USING NONRADIOACTIVE NUCLEIC ACID LABELING AND DETECTION SYSTEMS

Nonisotopically labeled probes have become increasingly popular in recent years and replace radioactive probes in techniques such as in situ hybridization, Southern, Northern and Western blot analysis. The advantages of these techniques are:

  • the technologies are safe
  • no hazardous radioactive waste accumulations
  • Probes can be stored for at least a year.
  • Hybridization solutions can be reused several times.
  • shorter exposure times

Types of nonradioactive labeling and detection systems used include the biotin-streptavidin and the digoxigenin –antidigoxigenin system.

In the biotin system biotin is incorporated in the probe by using biotinylated dNTPs during probe synthesis. The incorporated biotin is detected directly by avidin or streptavidin or an anti-biotin antibody conjugated to a fluorochrome or an enzyme such as alkaline phosphatase or horseradish peroxidase. Avidin is a 68 kD glycoprotein derived from egg white and streptavidin is a 60 kD protein from Streptomyces avidinii.

The digoxigenin-anti-digoxigenin system uses digoxigenin (DIG), a cardenolide steroid isolated from Digitalis plants.

In both systems the probe is detected with either chromogenic (colorimetric) substrates, fluorescence or chemiluminescence (see later).

4. THE DIGOXIGENIN SYSTEM

The DIG System is an effective system for the labeling and detection of DNA, RNA, and oligonucleotides. The protocols for labeling with digoxigenin (Figure 1) and subsequent detection are based on well established, widely used methods. DNA, RNA, and oligonucleotide probes are labeled according to the methods (usually enzymatic) used for preparing radioactive probes. Hybridization of digoxigenin-labeled probes (e.g., to target DNA or RNA on a Southern or Northern blot) is also carried out according to standard protocols, except that a special blocking reagent is used to eliminate background. The signal on the nucleic acid blot is detected according to the methods developed for western blots. The incorporation and spacing of digoxigenin in DNA, RNA, and oligonucleotides can be varied by using different labeling protocols.

4.1. DIG Labeling protocols

  • PCR Labeling: DIG probe synthesis incorporates Digoxigenin-11-dUTP by the polymerase chain reaction. (1 every 10–20 nucleotids)
  • Random-primed labeling method: Digoxigenin-11-dUTP is incorporated (1 every 20-25 nucleotides) by using the Klenow Polymerase and hexaoligonucleotides as primers.
  • T7/SP6-mediated transcription for the synthesis of strand-specific RNA probes. Using the RNA polymerases T7 or SP6 a DIG labeled RNA probe can be synthesis during transcription by incorporating Digoxigenin-11-UTP
  • 3'-End Labeling: Terminal transferase adds a single Digoxigenin-11-ddUTP to the 3'-end of the oligonucleotide.
  • DIG Oligonucleotide Tailing: Terminal transferase adds a string of Digoxigenin-11-dUTP interspersed with unlabeled dATP to the 3'-end of oligonucleotides.
  • DIG Oligonucleotide 5'-End Labeling: DIG-NHS ester labels the 5' end.
  • DIG labeling by nick translation: Uses DNA Polymerase I to create singel-stranded nicks in double stranded DNA. The 5'-3' exonuclease activity of E.coli Polymerase I enters the nicks and removes stretches of single-stranded DNA; the degraded DNA is than replaced by the 5'-3' polymerase activity.
  • cDNA synthesis: Using AMV (Avian Myelobastrosis Virus) reverse transcriptase the cDNA is labeled by incorporation of Digoxigenin-11-dUTP during cDNA synthesis.

The yield of the labeling reaction should be estimated to ensure the success of the reaction and to approximate the amount of probe to be used in the hybridization experiment. A simple dot blot method is used to estimate probe yield.

4.2. DIG Detection

Several alternatives are available for the detection of digoxigenin (or biotin)-labeled probes.

  • luminescent detection uses the chemiluminescent alkaline phosphatase substrate CSPD® to produce a light signal, which is detected by exposing the membrane to an X-ray film.

Figure 3: Schematic of the CSPD reaction: Enzymatic dephosphorilation of the dioxetane CSPD by alkaline phosphatase leads to the metastable phenolate anion, which decomposes and emits light at 477 nm.

  • colorimetric detection uses the substrates NBT and BCIP to generate purple/brown precipitate directly on the membrane.

Figure 4: Schematic of the NBT/BCIP reaction: when alkaline phosphatase removes the phosphate group of BCIP (5-bromo-4-chloro-3-indolyl-phosphate) the resulting molecules dimerizes under oxidating conditions to give the blue precipitate (5, 5'-dibromo-4,4'-dichloro-indigo). During the reaction with BCIP, NBT (nitroblue tetrazolium) is reduced to its colored form to give an enhanced color reaction.

  • multicolor detection comprises three naphthol-AS-phosphate/diazonium salt combinations for the visualization of a green, red, or blue hybridization signal.

Figure 5: Principle of the multicolor detection system: detection of the different labels is performed by binding the respective antibody- or streptavidine-alkaline phosphatase conjugate. The reactions are performed consecutively with heat-inactivationof the alkaline phosphatase between the detections. Hybrids must be stabilized by UV-crosslinking if mulitcolor detection is used.

  • There is also a wide range of alternative anti-digoxigenin conjugates available, such as anti-digoxigenin-peroxidase, anti-digoxigenin-gold (for electron microscopic studies), anti-digoxigenin-fluorescein and anti-digoxigein-rhodamine (for fluorescence detection in the microscope or with a fluorimeter).

4.3. General Considerations for Hybridization with DIG-probes

Please review this section of general hybridization considerations before proceeding with the DIG-system. Several points are critical for successful use of the DIG-system, especially when performing chemiluminescent detection.

4.3.1. Membrane Selection

For best results, use positively charged Nylon membranes for the transfer. Uncharged membranes can also be used with the DIG-system. Their binding capacity is lower and therefore a lower maximum sensitivity can be achieved.

Nitrocellulose membranes cannot be recommended in combination with the DIG-System. They can only be used when colorimetric detection will be performed and no stripping and reprobing is planned.

4.3.2. Probe Concentration

In the following chapters we give recommendations for probe concentrations in the different applications. These recommendations refer to newly synthesized, DIG-labeled probe. The labeling efficiency has always to be confirmed by estimating the yield of a labeling reaction. The recommended probe concentration must be regarded as a starting point for your hybridization.

Note: If chemiluminescent detection is performed, a too high probe concentration will often lead to background. Therefore, the probe concentration should not be increased above the recommended concentrations.

Figure 7: Mock hybridization and effect probe concentration. Naked pieces of membrane were incubated with the indicated amounts of DIG-labeled DNA probe and detected with chemilumininescence.

4.3.3. Optimization of the probe concentration – the “mock” hybridization

To prevent background problems as a result of a too high probe concentration, we recommend optimizing the probe concentration in a mock hybridization before the actual hybridization is performed.

The mock hybridization is carried out by incubating small membrane pieces (without DNA transferred to it) with different probe concentrations in the hybridization solution and subsequent detection with the procedure of choice.

For example, the highest probe concentration that gives an acceptable background should be used for the hybridization experiment (see figure 7, 25 ng/ml).

4.3.4. Hybridization and Washing Conditions

We have found that DIG-labeled probes demonstrate the same hybridization kinetics as radiolabeled probes. Hybridization and washing conditions for DIG-labeled probes do not differ substantially from those of radiolabeled probes.

The optimal hybridization and wash conditions for each probe must be determined experimentally. In this User’s Guide, we provide recommendations for hybridization and washing conditions. Use the conditions given as a starting point. It may then be necessary to optimize conditions to obtain maximum sensitivity with your probe.

Labeled probes can hybridize non-specifically to sequences that bear homology but are not entirely homologous to the probe sequence. Such hybrids are less stable than perfectly matched hybrids. They can be dissociated by performing washes of various stringencies. The stringency of washes can be manipulated by varying the salt concentration and temperature. For some applications, the stringency of the washes should be higher. However, we recommend that you hybridize stringently rather than wash stringently.

4.3.5. Prehybridization/Hybridization solutions

Several hybridization buffers can be used with the DIG-System. The main difference with hybridization buffers described elsewhere, is the presence of Blocking Reagent. The protein in Blocking Reagent reduces the non-specific binding of probe to the membrane filter.

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DIG Easy Hyb*

 

Standard buffer
+ 50% formamide

High SDS buffer
(Church buffer)

_________________________________________________________________________


ready-to-use
solution, non-
toxic used like
formamide-based
hybridization
solution

7% SDS, 50% formamide,
deionized, 5 x SSC, 2%
Blocking Reagent, 50 mM
sodium phosphate pH 7.0,
0.1% (w/v) N-lauroylsarcosine

50% formamide, deionized,
5 x SSC, 0.1% (w/v) N-lauroyl-
sarcosine, 0.02 (w/v) SDS,
2% Blocking Reagent

5 x SSC, 0.1% (w/v)
N-lauroylsarcosine,
0.02% (w/v) SDS,
1% Blocking Reagent


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4.3.6. Hybridization containers

You may use sealable containers or heat sealable plastic bags for the hybridization if using DIG Easy Hyb solutions. Roller tubes in combination with a hybridization oven may also be used for standard buffers.

4.3.7. Storage and Reuse of Hybridization Solutions

One of the advantages of the DIG-System is the stability of the labeled probe. After hybridization against the blotted target, the hybridization solution still contains large amounts of unannealed DIG-labeled probe. Simply pour the solution into a plastic tube and store at –20° C for DNA probes and –70°C for RNA probes. DIG labeled probes are stable for at least 1 year when stored in this manner. For reuse, thaw and denature by heating to +95°C for 10 min. If the hybridization solution contains formamide or if DIG Easy Hyb was used, denature at 68° C for 10 min.

4.3.8. Stripping and reprobing

With the DIG-System, membranes can be stripped and reprobed.

Note: When reprobing is planned, membranes must be kept wet at all stages after the first probe has been applied.

5. THE DIG SYSTEM AND SOUTHERN BLOTTING

The DIG-System can detect 0.03 pg (chemiluminescent detection) or 0.1 pg (colorimetric detection) homologous DNA in a Southern blot format on a nylon membrane. This corres-ponds to the detection of a single-copy gene in <1 µg of human genomic DNA. The proced-ures described here, are used routinely in our labs and have been found to give optimal results in Southern blotting, particularly in genomic Southern blotting.

5.1. Gel Electrophoresis

Restriction digest the DNA. Prepare an agarose gel of appropriate percentage, using a high-purity, nucleic acid grade agarose and Tris-Borate-EDTA (TBE)- or Tris-Acetate-EDTA (TAE)-buffer. Run the digest on the gel. If desired, the gel may be stained with ethidium bromide to visualize DNA fragments and to confirm the subsequent transfer to the membrane.

5.2. Southern Transfer

The transfer of DNA from the gel to the membrane can be accomplished by one of a number of common procedures; however the following procedures are routinely used in our lab and provide optimal detection sensitivity.

5.2.1. Depurination (optional)

Controlled acid treatment depurinates DNA. In the subsequent alkaline denaturation of the DNA, the DNA-strand breaks at the depurinated sites, resulting in smaller, easier to transfer fragments. Depurination is an optional treatment, usually performed when fragments >10 kb must be transferred. If you are transferring small DNAs (<10 kb) or detecting only the smaller fragments in a genomic digest, it may not be necessary to depurinate the DNA. Avoid excessive acid treatment; the fragments will be too small, which results in poor detection sensitivity.

Procedure

  1. Submerge the gel in 250 mM HCl for 10 min, with shaking, at room temperature. Do not exceed 10 min.
  2. Rinse the gel with H2O before proceeding to the “Denaturation section”.

5.2.2. Denaturation, neutralization, and blotting

  1. Submerge the gel in denaturation solution for 2 x 15 min at room temperature. Shake gently. This treatment denatures the DNA, making it single-stranded and accessible for the later applied probe.
  2. Rinse the gel with H2O.
  3. Submerge the gel in neutralization solution for 2 x 15 min at room temperature.
  4. Prepare membrane filters for Southern transfer. Nylon Membranes can be used without any prior treatments. Always use unpowdered rubber gloves when handling membranes, and manipulate the membranes with forceps at the edges only.
  5. Check pH. This is especially necessary when working with nitrocellulose. The pH should be <9, but nylon membranes also tolerate a higher pH. Especially when DNA transfer to nitrocellulose membranes is intended, it is important to check the actual pH of the gel after neutralization. It should be below pH 9 (nylon membranes will tolerate a higher pH), otherwise membranes will turn yellow and break during hybridization. To check the pH of the gel, lift one edge of the gel where no DNA has been loaded, press a pH stick into it and read the pH.
  6. Blot the DNA from the gel by capillary transfer to the membrane, using 20 x SSC buffer. Blot overnight to ensure efficient transfer of the DNA. Alternatively, the DNA can be vacuum-blotted onto the membrane; vacuum blotting can be accomplished in 1–2 h, according to the manufacturer’s recommendations. Our experience indicates that capillary transfer is more efficient than vacuum transfer.

5.2.3. Fixation

Crosslink the DNA to the membrane by any of the following procedures.

  • UV-crosslink the wet membrane without prior washing. After the UV crosslinking, rinse the membrane briefly in H2O and allow to air-dry. For UV-crosslinking of membranes special devices are available, that perform better than transilluminators.
  • Or bake the membrane at +120°C for 30 min
  • Nitrocellulose membranes must be baked at +80°C and under vacuum, to prevent spontaneous combustion of the nitrocellulose.
  • The membrane can now be used immediately for prehybridization, or can be stored dry at +4°C for future use.

5.2.4. Prehybridization and Hybridization

Prehybridization prepares the membrane for probe hybridization by blocking non-specific nucleic acid-binding sites on the membrane. This ultimately serves to lower background. Many different prehybridization solutions have been described in the literature. However, the prehybridization solutions described here combine efficient blocking with ease of use. As with any probe, optimal hybridization conditions for DIG-labeled probes must be determined experimentally. We strongly recommend that the time be taken to optimize each DIG-labeled probe (see the mock hybridization). The time taken for optimization will result in cleaner results and, ultimately, time savings, especially if a probe will be reused many times.

Table 7. Optimal hybridization conditions for different probes types. Optimal hybridization conditions for different probes types.

_________________________________________________________________________


Probe type

Temperature for prehybridization and hybridization*

Hybridization solution

Probe concentration

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DNA

 

 

 

 

Hybridize overnight at 37–42° C
Hybridize overnight at 65–68° C
Hybridize overnight at 37–42° C
 
Hybridize overnight at 37–42° C

DIG Easy Hyb
Standard buffer
Standard buffer
+ 50% formamide
High SDS buffer

5–25 ng/ml**

 

 

 

 

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RNA

 

 

Hybridize overnight at 50° C
Hybridize overnight at 50° C

 

DIG Easy Hyb
Standard buffer
+ 50% formamide

100 ng/ml**

 

 

_________________________________________________________________________


ready-to-use
solution, non-
toxic used like
formamide-based
hybridization
solution

7% SDS, 50% formamide,
deionized, 5 x SSC, 2%
Blocking Reagent, 50 mM
sodium phosphate pH 7.0,
0.1% (w/v) N-lauroylsarcosine

50% formamide, deionized,
5 x SSC, 0.1% (w/v) N-lauroyl-
sarcosine, 0.02 (w/v) SDS,
2% Blocking Reagent

5 x SSC, 0.1% (w/v)
N-lauroylsarcosine,
0.02% (w/v) SDS,
1% Blocking Reagent

_________________________________________________________________________



**The conditions given here are stringent conditions applicable if probe and target have 100% homology and a GC content of about 50%.

**When CDP- Staris used for detection the recommended concentrations are 10–20 ng/ml DIG-labeled DNA or 20–50 ng/ml DIG-labeled RNA. Higher concentrations may cause background.

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Procedure

  1. Place the blot in a hybridization bag containing 20 ml prehybridization solution per 100 cm2 of membrane surface area. Seal the bag, and prehybridize at the anticipated hybridization temperature for 2 h. Longer prehybridization times are acceptable. Several membranes can be processed in the same sealed bag as long as there is sufficient prehybridization solution to cover all the membranes, and the membranes can move freely in the bag. The optimal hybridization temperature for a specific probe will depend on the length of the probe and on the extent of sequence homology with the target sequence; therefore, it must be determined empirically. See Table 7 for recommended temperatures for different types of probes and different hybridization solutions.
  2. When using double-stranded DNA probes, heat in a boiling water bath for 10 min to denature the DNA. Chill directly on ice. Single-stranded RNA probes and oligo-nucleotide probes do not require denaturation prior to dilution unless extensive secondary structure is predicted from the sequence. Prepare at least 3.5 ml hybridization solution for a blot of 10 x 10 cm.
  3. Dilute the probe in hybridization solution. See Table 7 for optimal probe concentrations.
  4. Discard the prehybridization solution from the bag. Add the hybridization solution con-taining the DIG-labeled probe. Allow the probe to hybridize. See Table 7 for selecting a hybridization solution and temperature.
  5. At the end of the hybridization, pour the hybridization solution from the bag into a tube (with cap) that can withstand freezing and boiling (e.g., 50 ml polypropylene). Note: This hybridization solution contains unannealed DIG-labeled probe. The entire solution can be reused in future hybridization experiments. Label and date the tube, and store DNA probe solutions at –20°C and RNA probe solutions at –70°C. DIG-labeled probes stored in this manner are stable for at least 1 year. For reuse, thaw and denature by heating to +95° C for 10 min. If the hybridization solution contains 50% formamide (the flash point of pure formamide is +68°C) or DIG Easy Hyb, denature at +68° C for 10 min.
  6. Wash the membrane twice, 5 min per wash, in 2 x wash solution at room temperature. These washes (steps 6 and 7) remove unbound probe, which will lead to high backgrounds if not removed.
  7. 7. Wash the membrane twice, 15 min per wash, in 0.1 x wash solution. Long probes (>100 bp) should be washed at 68°C. For shorter probes, the wash temperature must be determined empirically.

Note: For most applications, washing in 0.5 x wash solution is stringent enough. It must be determined empirically whether it is necessary to wash with 0.1 x wash solution (0.1 x SSC, containing 0.1% SDS.

5.3. Detection of DIG-labeled Nucleic Acids

5.3.1. Chemiluminescent Detection

Using chemiluminescent detection, a light signal is produced on the site of the hybridized probe. The light signal can be recorded on X-ray films, requiring only very short exposure times. Chemiluminescent detection is a three-step process. In the first step, membranes are treated with Blocking Reagent to prevent nonspecific attraction of antibody to the membrane. Then, membranes are incubated with a dilution of Anti-Digoxigenin, Fab fragments, which are conjugated to alkaline phosphatase. In the third step, the membrane carrying the hybridized probe and bound antibody conjugate is reacted with a chemiluminescent substrate and exposed to X-ray film to record the chemiluminescent signal detection.

Chemiluninescent alkaline phosphatase substrates are available as CSPD ® or CDP-Star™ and their use depend on your experience in working with chemiluminescent detection. CDP-Star™ is the fastest chemiluminescent substrate available, typically requiring exposure times of only 15–60 s, whereas exposure times for CSPD ® are typically 15–30 min. Because working with CDP-Star™ requires some experience, we recommend to start your experiments with CSPD ®.

5.3.2. Guidelines for handling CSPD ® or CDP-Star™

To maintain full activity, as well as the nuclease-, phosphatase-, and bacteria-free environment in which the chemiluminescent substrates CSPD ® or CPD-Star™ are provided, adhere to the following precautions:

  • Do not place any non-sterile instrument (e.g., pipet tips) into the substrate solutions.
  • Remove the substrate from the bottle by pouring it into a sterile container using sterile technique or by transferring it with sterile pipettes. Wear powder-free gloves, and avoid touching the mouth of the bottle to anything.
  • Avoid touching the membrane with fingers (gloved or ungloved).
  • Use blunt-ended forceps that have been washed and autoclaved (to avoid alkaline phosphatase contamination) to pick up membranes, and handle membranes only at their edges.
  • Wear unpowdered gloves, and use hybridization bags free of dust and powder. Gloves or bags can be washed in distilled water before use.

Procedure

Perform all incubations at room temperature. Incubations can be performed in a sealed hybridization bag or clean plastic tray. If a bag is used, remove all large air bubbles that may be present in the bag. If a tray is used, agitate the tray gently to ensure that the membrane is always covered. If you are using more than one membrane, add enough solution to cover all membranes.

  1. After hybridization and post-hybridization washes, equilibrate the membrane in washing buffer for 1 min.
  2. Allow the chemiluminescent substrate to come to room temperature.
  3. Using a freshly washed dish or bag, block the membrane by gently agitating it in blocking solution for 30–60 min. Near the end of the blocking period, prepare the antibody solution as described in step 4. Longer blocking times are acceptable.
  4. Dilute the Anti-Digoxigenin-AP 1:10,000 in blocking solution. Small antibody aggregates in the Anti-Digoxigenin-AP may lead to background in the detection. It is therefore recommended to centrifuge the vial with antibody conjugate for 5 min at 13,000 rpm, before the first use. After the first use it is sufficient to centrifuge the Anti-Digoxigenin-AP for 1 min, directly before dilution. Mix gently by inversion. For example, for a 1:10,000 dilution, add 3 µl Anti-Digoxigenin-AP to 30 ml blocking solution and mix. When working with CDP-Star™, dilute Anti-Digoxigenin-AP 1: 20,000 in blocking solution. This working antibody solution is stable for about 12 h at +4° C.
  5. Pour off the blocking solution and incubate the membrane for 30 min in the antibody solution prepared in step 4.
  6. Discard the antibody solution. Gently wash the membrane twice, 15 min per wash, in washing buffer.
  7. Pour off washing buffer and equilibrate the membrane in detection buffer for 2 min. It is important that the filter is kept wet before the chemiluminescent substrate is applied. If the membrane is even slightly dry, high background can occur.
  8. There are two methods of applying the diluted substrate. The single-filter method should be used when DNA is to be visualized on a single membrane. The filter-batching method is recommended for multiple membranes but may also be used when visualization is performed on a single membrane.
  9. Single-filter method

    1. Place the membrane between two sheets of acetate (plastic page protectors). Gently lift the top sheet of plastic and, with a sterile pipet, add approximately 0.5 ml (per 100 cm2 ) of the chemiluminescent substrate on top of the membrane, scattering the drops over the surface of the membrane. Lower the top sheet of plastic. With a damp lab tissue gently wipe the top sheet to remove any bubbles present under the sheet and to create a liquid seal around the membrane. Incubate filter for 5 min. Proceed to step 9.

    Filter-batching method

    1. Pipette 5–10 ml of diluted CSPD ® or CDP-Star™ into the center of sterile dish. Using blunt-end forceps, place the membrane in the dish. Tilt the dish until the membrane is thoroughly saturated.
    2. Incubate filter for 5 min. Remove the membrane from the substrate, and allow any excess liquid to drip off. Do not allow the membrane to dry.
    3. Cover the damp membrane by placing it between two clear acetate sheets or page protectors.
    4. Wipe the top sheet with a damp lab tissue to remove any bubbles present between the sheet and the membrane.
    5. Repeat the filter-batching method until the chemiluminescent substrate has been applied to all membranes. To prevent the membrane from drying out, avoid repeated exposure to air. After treating the final membrane, proceed to step 9.
  10. Seal the semi-dry membranes in a plastic bag.
  11. For the briefest exposure to X-ray film, the alkaline phosphatase chemiluminescent reaction must be at a steady state. At room temperature, 7–8 h are required to reach a steady state reaction. Once a steady state is reached, single-copy gene detection on a human genomic blot can be obtained with an approximate exposure time of 15 min. If the membrane is exposed before the steady state is reached, approximately 60 min of exposure is required for single-copy gene detection on a human genomic blot. Therefore, to shorten exposure times, we recommend incubation of the membrane for 15 min at +37°C before exposure to X-ray film.
    1. Note: Because CDP-Star™ shows higher initial signals, this incubation step is neither necessary, nor recommended when CDP-Star™ is used.
  12. For detection of the chemiluminescent signal, the membrane is exposed to standard X-ray film. Multiple exposures from a single blot can be obtained for up to 2 days after the addition of the chemiluminescent substrate. As starting point for the exposure time, we recommend 15–20 min (CSPD ® ) or 1 min (CDP-Star™). Adjust the exposure time to the signal strength.

5.4. Stripping Membranes for Reprobing

Stripping of blots that have been detected colorimetrically is only possible when nylon membranes were used for blotting. The color precipitate has to be removed by incubation in dimethylformamide (DMF) and nitrocellulose is dissolved under such conditions.

The luminescent signal can be easily removed by a short wash of the filter in water. For the subsequent removal of probe there are several procedures. With the procedure described below we have achieved good results.

Note: When a membrane is to be rehybridized the membrane should be kept wet at all stages between hybridization and probe removal

Method I

  1. Heat 100 ml of 0.1% SDS in a 500 ml beaker.
  2. Shortly before the SDS-solution starts to boil, transfer the membrane to a clean tray.
  3. Pour the boiling SDS-solution over the membrane.
  4. Incubate for 10 min on a rocking platform (i.e., without further heating).
  5. Wash for 5 min in Washing buffer at room temperature.
  6. Proceed to the prehybridization.

Note: After stripping, start with the prehybridization or store the filter wet in 2 x SSC in a sealed plastic bag.