Nucleic Acid Hybridization

NUCLEIC ACID HYBRIDIZATION PRINCIPLES

"Blotting" means transferring one thing to another in one fell swoop, e.g. to blot some ink onto a paper.
In 1975 Edwin Southern proposed to hybridize nucleic acids immobilized on a solid support. Until then, the hybridization had been performed only in solution (two chains pairing with each other to give a double helix in a test tube). The solid support (nitrocellulose, at the time) allowed maintaining a permanent copy of the molecules "recorded" in the position they had taken in the electrophoretic run, following the transfer of the molecules in the gel to the piece of nitrocellulose.
In order to identify specific DNA fragments, it had to be broken up.
DNA extraction and digestion, gel separation of the fragments and their transfer on a solid support, hybridization with a complementary probe to the specific sequence of interest are the main steps of the method. The stability of the bond between the probe and the target molecules depends on the percentage of similarity.
Hybridization can happen by base-pairing between any type of nucleic acid strand: DNA-DNA, hybrid DNA-RNA, RNA-RNA.

SOUTHERN BLOT FOR DNA

1. DNA enzymatic digestion. If we transfer undigested DNA, we would have molecules all of the same length, and therefore a single band would form. It is useful in this case to use restriction endonucleases able to cut at the inside of a DNA molecule. DNase is an aspecific endonuclease. "Restriction" means that the enzyme cuts at specific points. They have been detected in bacteria. After extraction, we have DNA fragments about 20-50 kb long. The restriction enzyme must have a cut-off frequency lower than the size of the molecules. So-called frequent cutters are used, they are enzymes that make frequent cuts on average in the genome. EcoRI recognizes a sequence of 6 bases.
We need:
- Purified and concentrated DNA substrate. If not purified, it may contain restriction enzyme inhibitors.
- An active enzyme.
- Optimal reaction conditions.
- Buffer with optimal saline concentration.
One Unit of the restriction enzyme is meant to be the amount of enzyme that digests 1 μg of DNA in one hour.
An over-digestion is usually desired (by increasing the reaction time or the amount of enzyme present) which ensures that all the sites are cut.
Class II enzymes are used (those recognizing shorter 4 base sequences cut too much). Restriction mixtures are generally made to have an optimal cut-off frequency. Not1 is a rare-cutter, recognizing a sequence of 8 bp.
Southern blot tells us if there is a gene, and how many copies there are compared to normal control.
Enzymes must be diluted very carefully because protein molecules in water tend to undergo denaturation. All enzymes are mixed with BSA (Bovine Serum Albumin), which is a stabilizer, and it does not allow the solution to be too diluted. They are stored at -20°C with 50% glycerol. Freezing damages the protein structure, because ice crystals break the protein molecule. Glycerol has a lower freezing point than water. At -20°C, the enzyme is more stable, while glycerol does not freeze.
The glycerol, in the final solution, must be at most 5%, an excess of it can give rise to phenomena such as "star activity": the enzyme can cut sequences that are not its specific target. If the digestion time increases, the amount of enzyme can be reduced. The enzyme is a catalyst, it is not lost or consumed. The functional efficacy of the enzyme can be checked during the work with a minigel.
Once the reaction has taken place, a standard agarose gel is made (1% in general) and a streak of bands is obtained.

The probe binds to all the complementary pieces of DNA that are on the gel, so more bands will be highlighted. When we have the truly complete DNA sequence, with "restriction simulators" (they are based on word processors: they can find sequence "text") it is possible to predict how a given gene will be cut by a particular mixture of restriction enzymes, except for polymorphisms.
There are restriction enzymes that cut the two DNA strands symmetrically (generating "blunt" DNA ends) or asymmetrically (generating "sticky" ends, with one strand protruding over the other).

2. Agarose gel electrophoresis of the DNA fragments.

3. Transfer (blotting) of the separated DNA fragments from the gel to a solid support (a sheet filter of nitrocellulose, or better of nylon, which is more robust and may be electrically (positively) charged to better retain DNA.).
a) The classic blot is a capillary blot: sheets of paper soaked in SSC (sodium saline citrate, a very concentrated solution of NaCl and Na citrate) attract the gel water that drags up the molecules that remain trapped in the filter that has molecular (sized) pores. The molecules rise in a straight line and get trapped in the same position they occupy in the gel.
If you don't stack everything well, it may fail.
b) Reverse blot: the gel is placed on the membrane, in this way, gravity is also exploited, and the saline solution is continuously dispensed from a wet sponge at the top of the blot stack.

In order to keep the DNA molecules denatured, so that they can be able to hybridize with the probe, the blotted filter can be soaked in an alkaline solution, or blot can be directly performed using NaOH 0.4 M as the transfer medium.

4. Fixation of the DNA to the filter. By using a UV lamp at 254 nm, at a certain distance from the filter and for 5 minutes, the chemical groups of sugar and PO₄ are energized so that they bind to the amino groups -NH2 of the nylon membrane. Our standard exposure is 12 watts, about 15 cm, for 5 minutes. Following fixation, the blotted membrane can be hybridized up to 12 times in our hands. If it is fixed too much, however, due to the formation of too many bonds with the filter, the bases are made unavailable for hybridization. The same is also obtained by providing temperature: putting the filter in the microwave for a few minutes.
A nylon sheet is obtained with all the lanes and here the fragments of DNA separated by size.

5. Probe labelling

A) Random priming
The easiest way is to use an in vitro polymerase that copies the filaments using a radioactive nucleotide: ³²P dCTP.
In the test tube: cDNA probe, random primers, in order to prime synthesis of the labelled strand from any template sequence, DNA polymerase, the 4 dNTPs, one of which labelled in one of the three phosphate groups, which are called according to their proximity to the sugar from alpha to gamma:

base-sugar-P-P-P
α β γ
An α-labelled ³²P is purchased because, in the polymerization reaction, only alpha P remains in the synthesized strand. It must be kept in mind that "alpha" does not indicate here the type of emission of ³²P, which is a β-emitter (fast electrons), but the position of the radioactive P.

B) Oligo terminal (3´ ) labelling
If a ready-made single-strand oligonucleotide is the probe to be labelled, it will not have the 5´phosphate that natural molecules have, instead, it has a classic 3´-hydroxyl end. The terminal deoxynucleotidyl transferase (TdT) is used in this case (recombinant TDT). This enzyme transfers deoxynucleotides to the terminal position at the extreme 3´. It is an example of a polymerase (an enzyme that extends a DNA polymer by catalyzing the phosphodiester bond) that does not need a template. It is found only in pre-lymphocytes (a marker for the diagnosis of leukaemia) and works in the presence of divalent cations: if the added nucleotide is a purine, it is better to use Mg⁺⁺, if it is a pyrimidine it is better to use Co⁺⁺. Single nucleotides can be added using deoxynucleotides. In this case, in the reaction tube we put:
- the oligonucleotide to be marked,
- dATP where the alpha P is a beta-emitting ³²P (ATP because it seems that the TdT has a greater affinity for the A),
- the TdT.
A chain of radioactive As will be added at the 3 ´ end of the oligo molecules.

C) Oligo 5´ labelling
By using a gamma ³²P deoxynucleotide and the enzyme PNK (Polynucleotide Kinase), the gamma phosphate may be transferred to the 5´ end of a single strand probe whose original 5´ phosphate group has been removed by alkaline phosphatase. In this way, a single radioactive atom is added to each probe molecule, so that this is a weaker labelling.

6. Hybridization. Hybridization consists in making the filter react with a probe to verify the existence, quantity and possible alterations of a particular fragment of DNA.
Steps:
- Probe preparation (in solution). The probe must be heated at 94-100°C to denature it before adding it to the hybridization reaction.
- Renaturation kinetics: the double helix is reformed between the probe and the target. There is the mixing of two single strands that make a double helix that was not there before. It is an indirect mode, the probe must be known and must also be visible (for us or for the detection instrument) in order to reveal an unknown and invisible molecule.

The hydrogen bonding between the pairs involves association and dissociation kinetics; stability increases with increasing lengths of complementary nucleic acid chains, which will favor association.
Many other factors can affect this equilibrium. The primary influences are concentration, temperature, and salt concentration in the hybridization buffer (Davis et al., 1986).
A molar excess of a nucleic acid molecule molecule will favor its pairing eith the complementary chain.

In the presence of adequate salt concentration (cations, to neutralize negatively charged DNA molecules allowing their pairing) and temperature conditions, only specific binding of the probe to the target will be favoured.

The stability of the double-stranded hybridized molecule is also affected by:
- the number of mismatches in the two paired strands;
- the percent of G-C bonds versus A-T bonds (a G-C bond involves three hydrogen bonds and two in A-T bonds):
- the amount of formamide in the buffer (formamide is a commonly used denaturing agent for DNA, influencing DNA duplex stability).

The incubation temperature (Τ_i) can be estimated from the following formulas:
Τ_i =T_m - 15°C
T_m = 16.6 log[M] + 0.41 [P_GC] + 81.5 - P_m - B/L - 0.65[P_f]

Where:
[M] is the molar concentration of Na⁺, to a maximum of 0.5 (1 x SSC contains 0.165 Μ Na⁺)
[P_GC] is the percent of G or C bases in the oligonucleotide probe (between 30 and 70)
P_m is the percent of mismatched bases, if known
(each percent of mismatch will alter the T_m by 1°C on the average)
Β is 675 (for synthetic probes up to 100 bases)
L is the probe length in bases
[P_f] is the percent of formamide in the buffer

16.6 log 0.165 + (0.41*50) + 81.5 - Pm=0 - 675/25 - (0.65*0)
16.6 * -0.78 = -12.99
- 12.99 + 20.5 + 81.5 - Pm=0 - 27 - 0 = 62°C

Under "stringent" hybridization conditions, one could also distinguish between target sequences divergent only for one base.
If "permissive" conditions are chosen, similar genes will be identified (members of gene families), slightly different from the known probe, also across different species. DNA and RNA have extraordinary flexibility to make single helix sections alternating with double helix sections.
Thus, any molecule that has a certain homology with the probe can be recognized, even if for example there are some blocked bases of the target.
Cocktails of oligonucleotides may be used to hybridize with different exons of the same gene.

7. Washes. Removal of all that is weakly bound. In the classic case, we use a probe as specific as possible.
Wash stringency may be adjusted by rising temperature and/or lowering salt concentration: both these conditions tend to detach any nonspecifically bound probe molecules from the hybridized filter.
As an extreme case, when all the probe must be removed in order to start a new hybridization experiment with a different, the blot membrane is boiled in distilled water (the highest temperature, in absence of salts).

8. Autoradiography. It highlights the base pairing between the probe and the target. A photosensitive film covered with a photographic (radiographic) emulsion made of silver halide is used. The beta particles emitted by the labelled probe hit the electrons of this substance and cause the silver to release and form a precipitate, a black dot that we can see on the developed film. ³²P emits beta particles with high energy content. The hybridized, radioactive filter is wrapped in a plastic sheet to avoid contamination. The silver precipitate is directly proportional to the number of beta particles emitted; however, there is a slight widening of the bands, a blur around the point of origin, this phenomenon can be a problem for nearby bands that can overlap. To improve autoradiography, we can:
1) add an intensifying screen. It is screen coated with calcium tungstate. The most energetic beta particles that manage to cross the film hit the screen which covers the inside of the cassette, this generate new particles that come back and hit the film again. Sensitivity increases but the blur increases.
2) place the autoradiographic cassette in the freezer at -20°C (or even in the freezer -at 80°C). The reaction that occurs in a radiography is the precipitation of silver, and any precipitation is favoured at low temperatures.

NORTHERN BLOT FOR RNA

It is useful for studying the expression of a gene: in what tissues and periods of life is it active, how active is it, etc.

The general procedure in similar to Southern Blot, with some variation that will be highlighted.

Ribosomal RNA (rRNA) may be a useful reference point.
If there is degradation, the 28S tends to break into two pieces that end in the 18S-band, in this case, the brilliance of the 18S-band increases and that of the 28S decreases.
When working with rRNA to obtain a good resolution, a maxigel is made with larger wells to be able to load more RNA in order to have an appreciable signal (10 to 20 μg). The average quantity of RNA per cell is about 10 pg; therefore, it is necessary to start from about 1 x 10⁶ cells. From 10 mL of blood, about 10 μg of RNA are obtained; from other tissues, it is easier to have larger quantities.

RNA is run on the gel as much as possible to separate the bands well. Formaldehyde is added in the gel to inhibit the formation of hydrogen bonds. If it is not used, single-stranded filaments tend to fold due to self-complementarity. RNA has a very strong tendency to make double helices in complementary regions.
The Northern gel is blotted (in neutral citrated saline solution because the denaturant is already there: it is formaldehyde) on a nylon membrane. Ethidium bromide is also loaded into the well to be able to highlight the 28S and 18S bands for UV brilliance. A mark can be made by a pencil in correspondence the centre of the two bands, thus obtaining two points corresponding to molecules of known length and allowing building a straight line on which to evaluate the other bands.

There is a second possibility which consists in separating the polyadenylated messengers after removing the ribosomal RNA. On the filter only the putative messengers remain, the problem is that if there is also a trace of ribosomal RNA, there is the risk of having an unspecific signal for a cross-hybridization (cross-hybridization), even a low homology is enough because many copies of ribosomal not eliminated give a signal (bright band). For marker, a known marker is used. The rRNA release procedure also reduces the number of molecules that interest us: at least 50-100 μg of starting is required (e.g. 4 flasks of cells). We can hybridize the membrane as for Southern and then do autoradiography. We can expect to see even completely white film if the cells do not express that messenger, for this reason, some control points are needed: positive controls: at least one known RNA (e.g. 2 Kb) certainly present in the sample, indicates that the method has worked.

INTERPRETATION OF NORTHERN DATA

Reference markers are used: "housekeeping" genes (the housekeepers, who manage the house) are not specific to certain cells, they are active in all cells, and have two fundamental characteristics for their use in this sense:
1) are always expressed;
2) have a constant, and high, level of expression.
Northern manages to see RNA up to 5-10 copies per cell. The housekeeping gene must be expressed far more than 5-10 copies per cell.
The most used genes are:
- BETA-ACTIN (ACTB) mRNA (cytoskeleton).
- β2 MICROGLOBULIN (B2M) is expressed in all nucleated cells.
- GLUCOSE-3-PHOSPHATE-DEHYDROGENASE (G3PDH),
encoding for an enzyme involved in basic carbohydrate metabolism.
Housekeeping genes can be used in a Northern quantity to calibrate the expression of the gene on the quantity, for example of the actin band. These genes have also been found to have variations in expression as a function of the cell cycle phase. Β2M appears to be less affected by the cell cycle. However, it is less expressed.
By using Northern blot, kinetic studies can be done: at what time after stimulation is the peak of mRNA.

Additional bands occurring besides the mRNA expected band could be due to:

1. Primary, immature transcripts.
They are generally processed quickly, however sometimes it can be seen. It has many regions that are not there in the mature (intronic regions), and it can be seen in the intermediate stage of processing (the processing kinetics is very different).
2. Alternative splicing RNA isoforms,
also hybridizing to the probe.
3. Member of the same gene family,
whose mRNAs partly hybridize with a probe recognizing a similar member of the same family.
4. Cross-hybridization
with 28S or 18S RNA, seen in all all lanes if there is partial homology with these molecules very abundant in total RNA (85%).

An example of a Northern blot to characterize a newly discovered gene is here.

If the size of the expected mRNA is known a priori, and these size are very different, a filter can also be hybridized with several probes simultaneously.
Some vendors sell ready-made blots with 12 different human tissues, ready to be hybridized with a probe.