Ribosomal RNA Operon Copy Number Determination Via Southern Hybridization Using Non-Radioactive Detection

There are 7 distinct steps in this protocol: (elapsed time: 2 – 3 days)

  1. isolation of genomic DNA from the organism of interest
  2. digestion of genomic DNA with restriction endonucleases
  3. separation of digested genomic DNA via agarose gel electrophoresis.
  4. transfer & immobilization of gel-separated genomic DNA to a (+)-charged nylon membrane
  5. hybridization of digoxigenin(DIG)-labeled 16S rDNA probe to membrane-bound genomic DNA
  6. immunological detection of specifically bound probe with an alkaline phosphate-conjugated antibody specific to the DIG moiety on the DNA probe.
  7. detection of hybridizing DNA bands with an alkaline-phosphatase activated chemiluminescent substrate.

16S rDNA Probe Preparation (12 – 17 hr)

The probe used in hybridization experiments is prepared by PCR-amplification of a fragment of the 16S rRNA gene from Escherichia coli B/r genomic DNA using standard conditions and cycle times appropriate for your thermocycler. The PCR product is gel purified and labeled with digoxigenin(DIG)-dUTP using the DIG-High Prime random labeling kit following the manufacturer’s instructions (Roche Molecular Biochemicals). Experiments confirm that this conserved fragment and set of hybridization conditions are sufficient to accurately detect 16S rRNA genes from a phylogenetically diverse collection of prokaryotes.

PCR Primers

8f (5’-AGAGTTTGATCCTGGCTCAG-3’)

519r (5’-GTATTACCGCGGCTGCTGG-3’)

PCR Conditions

The probe is reusable, but signal intensity will decrease after each application. It is best to prepare excess probe and use fresh probe once you begin to see a reduction in signal (typically 5 – 10 uses). A 100 L PCR reaction will typically yield 1 – 4 g of product. I usually prepare four different 100 L reactions and pool them prior to gel purification of the fragment. Since thermocyclers vary with respect to temperature ramping times, use the conditions appropriate for your themocycler. Use a 60C annealing temperature with this probe combination to ensure that you only get one discrete band on your gel.

Gel Purfication of PCR Product (Probe)

Pool your PCR reactions and run out on a 1.5% agarose gel. Excise the fragment and gel purify with your method of choice. The QIAquick Gel Extraction Kit (Qiagen) is fast and produces good yields for gel purification of DNA fragments. Other protocols/kits are suitable as well. Keep in mind that most spin-filter methods are only capable of handling a limited quantity of DNA, so you may have to use more than one filter to purify your fragment.

Labeling of Gel Purified Fragment

The purified fragment is labeled with digoxigenin (DIG) dUTP using the DIG-High Prime Random Labeling and Detection Starter Kit II (Catalog No. 1 585 614) sold by Roche Molecular Biochemicals (formally Boehringer Mannheim). Follow the manufacturer’s instructions for labeling.

You will use 15 mL of a 25 ng/mL solution of labeled probe for hybridization. Therefore you should label at least 1 g of probe to obtain a sufficient amount of probe for a blot (15 mL x 25 ng/mL = 375 ng). I usually label 3 g of probe and store what is not used at -20C. Remember that the yield of the labeling reaction will not be 100%; therefore, if you label 325 ng of probe (quantified by A260nm) you will not obtain 325 ng of labeled probe. Th kit contains test strips and controls necessary to quantify the probe and calculate labeling efficiency.

To prepare in advance for hybridization, suspend 325 ng of labeled probe in 15 mL of hybridization buffer (recipe below) and store at -20C.

Restriction Digestion of Genomic DNA (2 hr – overnight)

It is important to perform several independent restriction digests in order to obtain an accurate estimate of the number of rRNA operons per genome. Two different situations during restriction digestion could lead to misleading results: 1) restriction digestion of 16S rRNA genes within the region targeted by the probe, leading to an overestimate of rRNA operon copy number, and 2) restriction digestion such that two or more 16S rRNA genes are on the same fragment, leading to an under-estimation of rRNA operon copy number. Restriction digestion is the critical step in rRNA operon copy number determination, therefore it is important to use high-quality enzymes and proper reaction conditions. I have been consistently disappointed with Gibco/BRL enzymes (incomplete digestion, low activity) and use only New England Biolab’s brand enzymes, which produce good results every time. When calculating reaction volumes, remember that restriction enzymes typically come in a 50% glycerol solution that must be diluted at least to 5% for optimum enzyme activity. The following enzymes have been used with success:

PstI, PvuII, SacI, AccI, PshA1, AvrII, RsrII, BstEII, SmaI, EcoRI.

Perform a minimum of six different restriction digests for each strain (one enzyme per digest, 1 – 2 g of genomic DNA per digest). Determine the number of rRNA operons from a minimum of 4 different digests producing identical hybridization results (equal number of bands of same intensity). When restriction digestion fails to resolve hybridized fragments or when bands of equal intensity cannot be discriminated, results should be discarded and additional analyses performed with different restriction endonucleases.

Gel Electrophoresis of Restriction Digested Genomic DNA (4 hr)

It is important to achieve maximum separation of restriction fragments with good resolution. I use a 12 cm wide x 14 cm long gel (3 – 5 mm thick) and run it at 85 volts for about 4 hr, at which time the bromophenol blue loading dye band (runs at around 700 bp) is at the end of the gel (0.75% agarose, 0.5X TBE buffer). DO NOT run the gel with stain in it (ethidium bromide retards migration of DNA and will give you smeared bands). There is no need to visualize the gel, but if you want to see the smear of DNA in each lane, post-stain the gel after running it, photograph, and then destain prior to Southern transfer. Begin transfer of DNA from gel to membrane immediately after gel run has ended to prevent diffusion of DNA from gel into running buffer.

Transfer & Immobilization of DNA from Gel to Nylon Membrane (2.5 hr)*

There are three different ways to transfer DNA from agarose gels to membranes: 1) capillary transfer, 2) electrophoretic transfer, and 3) vacuum transfer. The vacuum transfer protocol performed in our lab using the Pharmcia VacuGene XL vacuum blotter apparatus is described below.

Membrane Selection

Any positively charged nylon membrane should be suitable. We have used Hybond-N+ by Amersham, MagnaCharge by MSI (now Osmonics, Inc.), and Boehringer Manneheim’s (BMB) membranes with success (now Roche Molecular Biochemicals). Note that the MagnaCharge membrane will turn orange in color during the depurination step, but that does not affect the results.

Transfer Protocol

The transfer process involves placing the gel on the vacuum blotter and then pulling several solutions through the gel (and membrane below) with the applied vacuum.

Steps in Transfer Process:

  1. As gel nears end of run, assemble the vacuum blotter apparatus and cut a piece of nylon membrane approx. 1 cm. larger in width and length than the gel. Use gloves at all times when handling the membrane. Use forceps on edge of membrane when manipulating. Mark one corner for orientation with a soft lead pencil and record your mark for later identification.
  1. Wet the membrane for a few minutes in sterile distilled water (Milli-Q) prior to setting on vacuum support.
  1. Place wet membrane on the vacuum support, align the vacuum mask to “frame” the membrane on the support, and carefully place the gel on top (see diagram above). Turn on the vacuum pump and make sure the vacuum is pulling 40 mbar (higher pressure will collapse the gel matrix and prevent transfer of DNA to membrane). If you cannot get a vacuum, make sure that the gel is forming a seal by covering the mask completely. Avoid air bubbles between gel and membrane.
  1. Next you will apply a series of different solutions directly to the surface of the gel that are pulled through by the applied vacuum. (Note: the steps preceding transfer can be performed in a tray flooded with each solution in which the gel is submerged). The vacuum pressure will cause the gel to form a slightly concave surface that will keep the solutions on top of the gel. During each step, make sure that each solution completely covers the top of the gel. The length of time for the following steps is dependant upon the thickness of the agarose gel. The times below are appropriate for the 5 mm gel described above.
  1. Depurination (0.2 M HCl) – 20 min.

Note:this step is only necessary if you have DNA fragments >10 – 15 kb in your gel. Larger fragments do not transfer out of the gel as efficiently as smaller fragments of DNA. During this step the DNA is depurinated by acid treatment. Depurination is the chemical step where the purine base (adenine & guanine) is removed from the nucleotide moiety. DNA is susceptible to alkaki hydrolysis at depurinated sites, which fragments the DNA (next step).

  1. Denaturation (0.5 M NaOH, 1.5 M NaCl) – 20 min.

Two chemical processes are achieved in this step: 1) treatment of the gel in alkali solution hydrolyzes DNA at depurinated sites resulting in fragmentation of the DNA polymer, and 2) dsDNA is denatured into ssDNA under alkali conditions that enables binding of the DNA to the positively-charged nylon membrane. The alkali solution denatures DNA by disrupting the hydrogen bonds that hold the two strands of DNA together.

  1. Neutralization (1.0 M Tris, 1.5 M NaCl, pH 7.5) – 20 min.

Note: this step is not necessary for nylon membranes since they retain their binding capacity in alkaline solutions. The gel must be neutralized since base is not compatible with the nitrocellulose, and the charges on the DNA must be neutralized. Double stranded and charged DNA does not bind to nitrocellulose membranes.

  1. Transfer (20X SSC) 1.5 – 2.0 hr (thicker gels = longer transfer times)

During this step the DNA is transferred from the gel to the surface of the nylon membrane. Transfer is mediated by displacement of the DNA from the gel by the high ionic concentration of the transfer buffer. The vacuum pressure facilitates the rapid directional transfer of the DNA to the nylon membrane.

Note: Our laboratory has followed the method above, which consistently produces good results. Much of the above protocol is structured for use of nitrocellulose membranes that were initially the only membrane available for Southern hybridization. A much shorter procedure can be used with nylon membranes, but we have not tested this protocol as of yet. If it ain’t broke… The alternative method is called “Alkaline Transfer” and involves simply incubating the gel in excess Denaturation Solution for 15 min (twice with fresh solution), followed by vacuum transfer for 1 – 2 hr. Larger DNA (>10 – 15 kb) molecules can be fragmented by adding an initial depurination step as described above. After Transfer and UV Immobilization (see below) the membrane is incubated in Neutralization Solution for 5 min. to avoid altering the composition of buffers in subsequent steps.

  1. Turn off the vacuum and disassemble the vacuum blotter apparatus. Carefully place the membrane on a piece of Whatman filter paper and place in UV crosslinker.
  1. Immobilize the DNA to the nylon membrane by crosslinking with 1200 mJ of UV energy. UV radiation results in covalent binding of thymine bases (and other nucleotides to a lesser extent) and NH2 groups on the membrane surface (nitrocellulose) and other positively charged groups (proprietary information) on nylon membranes. You can “over-crosslink” your membrane – more is not better! The binding of DNA to the membrane is dose-dependent; overexposure to UV will result in complete binding of the DNA to the membrane and decreased hybridization capacity. Total exposure should be 120 mJ/cm2 for damp membranes.
  2. Carefully roll the membrane (around a sterile pipette works well) and place it in a hybridization bottle such that no part of the membrane (with target DNA) overlaps. It’s alright if the edges overlap since they do not contain DNA. You can trim off the edges without DNA, but I prefer to leave them to grab with the forceps without damaging the region to be probed. A hybridization bag is also a suitable alternative if you have a shaking water bath in which to incubate it.

Hybridization of Probe to Membrane-bound DNA (24 – 48 hr)

Three main steps are involved in hybridizing the probe to the membrane-bound DNA: 1) prehybridization, 2) hybridization, and 3) post-hybridization washing.

  1. Prehybridization
  2. Prewarm the Hybridization Solution to 45C in the hybridization oven, the temperature at which you will conduct the subsequent hybridization.
  1. Add 0.5 mL Hybridization Solution per cm2 membrane to the hybridization bottle (or bag) containing the membrane.
  1. Incubate 4 hr – overnight (This step can go over a weekend with no adverse effects). During this step the membrane is incubated in Hybridization Solution which contains several different chemicals that block non-specific sites on the membrane that are capable of binding free probe. The blocking compound in the Hybridization Solution is simply a highly purified form of casein digest (basically expensive powdered milk, which may work just fine, but has not been attempted in our lab). In addition, the Hybridization Solution contains SDS that also inhibits background binding and is an RNase inhibitor. The Na+ salts in the Hyb. Solution stabilizes DNA duplexes by neutralizing the negative charge of the phosphate backbone; the higher the salt concentration, the lower the stringency of the hybridization. Formamide in the Hybridization Solution relaxes secondary structure of DNA and is used to lower the effective Tm of your probe binding to target; generally, each percentage of formamide in the Hyb. Solution lowers the Tm of the probe by 0.4C.
  1. Hybridization
  2. Prepare your probe (best done day in advance) - 25 ng/mL probe in 15 ml Hybridization Solution. (12 hr – 24 hr).
  1. Denature the probe by placing your probe (in Hybridization Solution) in a 70C water bath for 15 min, followed by rapid cooling on ice. I usually make the probe up in a 50 mL conical tube and simply stick it in a bucket of ice to cool (to 40 - 50C).
  1. Discard the Hybridization Solution used from prehybidization step.
  1. Add the Hybridization Solution containing your probe to the bottle and place back in the incubator.
  1. Incubate at 45C for 12 hr – 24 hr. (This step can be run over the weekend with no adverse effects).
  1. Post-Hybridization Washing
  2. At the end of the hybridization, be sure NOT to discard your probe, it can be reused many times. Pour the Hybridization Solution containing your probe into a 50 mL conical tube and store at -20C for later use.
  1. Wash twice for 5 min. at room temperature in Washing Buffer I by filling the hybridization bottle with approx. 50 mL of buffer. This step is non-stringent and removes excess probe from the membrane
  1. Wash twice for 15 min. at room temperature in Washing Buffer II by filling the hybridization bottle with approx. 50 mL of buffer. This step is the stringent wash step that is used to reduce background and non-specific binding of the probe. Increased wash temperature and decreased salt concentration may be necessary to increase the stringency of this step.

Detection of Specifically Bound Probe (3 – 4 hr)

During this step an alkaline phosphatase conjugated antibody, specific to the digoxigenin-dUTP incorporated into your probe, is applied to the membrane, which activates a chemiluminescent substrate (CSPDTM) at the site of binding. See diagram below:

Steps to Detection

  1. Discard the Washing Buffer II from the previous step.
  1. Incubate the membrane in Blocking Solution for 30 min. at room temperature. During this step the non-specific antibody binding sites are blocked.
  1. Close to end of step 2, dilute the anti-DIG-AP conjugate 1:20,000 in Blocking Solution. To do so, add 1 L in 20 mL of Blocking Solution – this is the Antibody Solution.
  1. Discard the Blocking Solution from step 2 and add the Antibody Solution.
  1. Incubate the membrane for 30 min. at room temperature in Antibody Solution. During this step the antibody binds to the digoxigenin moiety incorporated in the probe as DIG-dUTP.
  1. Close to the end of step 5, fill a clean (!) tray with excess Washing Buffer III sufficient to cover your membrane.
  1. Discard the Antibody Solution, remove your membrane from the hybridization bottle and place it in the tray containing Washing Buffer III.
  1. Wash the membrane by placing the tray containing Washing Buffer III and your membrane on a rotating (or oscillating) platform shaker for 15 min. During this step you are removing excess, unbound, antibody that could otherwise produce non-specific signal.
  1. Prepare 15 – 25 ml of Detection Solution while your membrane is washing.
  1. Repeat step 8 with fresh Washing Buffer III.
  1. Carefully discard the Washing Buffer III and incubate your membrane in Detection Solution for 5 min. It is critical NOT to let this step exceed 5 min. During this step you are adding a buffer that establishes the optimum conditions for the alkaline phosphatase enzyme that is conjugated to the antibody, which is now bound to the probe, which is in turn bound to the target sequence.
  1. Place the membrane with the DNA side facing up on a sheet of overhead transparency film cut to fit in your x-ray film cassette (8 x 10 in). WORK FAST – DO NOT LET THE MEMBRANE DRY OUT!!! Dry membrane = high background. (Don't use transparency film for printers - the coating will bind the chemiluminescent substrate.)
  1. Apply 0.75 - 1.5 mL of CSPD (ready-to-use) to the damp membrane by carefully dropping the solution to cover the entire membrane. (Gently rolling a second piece of film over the membrane will help distribute the solution evenly.) CSPD is a luminescent substrate that is activated (cleavage of a protecting group) by alkaline phosphatase bound to the antibody.
  1. Place another sheet of transparency film on top to make a “membrane sandwich” and use a pipette to evenly distribute the CSPD solution back and forth across the membrane. Allow this set-up to incubate for 5 min. at room temperature on the bench-topj. DO NOT squeegee the solution yet.
  1. Using a glass pipette, carefully squeegee out excess CSPD solution without creating any bubbles.
  1. Dry off the exterior of the membrane sandwich with a tissue and place it on top of a sheet of Saran Wrap that is 1 – 2 cm larger in width and length than the transparency film. Wrap the protruding edges of the Saran Wrap around the membrane sandwich to seal the edges, preventing the membrane from drying out during incubation. Do not cover the DNA side of the membrane sandwich with Saran Wrap; this side should only be covered on the edges. See diagram below: