A native PAGE (polyacrylamide gel electrophoresis) was chosen to verify that our complementary sequences were forming the probe and to demonstrate that our miDNA can bind to the probe. We used a 12 percentage gel, and the polyacrylamide solution used was in a 1:19 ratio of acrylamide to bisacrylamide. The 12 percentage gel was able to distinguish a 22 nucleotide difference in the sample.
The native PAGE ensures that the base pairing between the complementary strands remains intact during the electrophoresis, which is required to verify the formation of our probe. The experiment was performed using standard SDS PAGE equipment, but a longer gel is ideal.
The zeta potential analyzer is used to find if the nanoparticles were evenly distributed in the solution and to confirm the presence of stable carboxylic acid groups. This system is designed for the characterization of colloidal, nanoparticulate, and macromolecular samples. It can determine the zeta potential, which is a measure of the electrostatic potential that exists at the interface between a particle and the surrounding liquid or dispersion medium.
The instrument works on the basis of Dynamic Light Scattering (DLS). It will detect the fluctuation in the intensity of scattered light over a time period, which is caused by the Brownian movement of particles and can calculate the zeta potential with this reading.
The experiment requires the samples to be diluted and loaded into a cuvette
The cuvette should be filled with around 500 microlitres of the aqueous solution of the particle. The cuvette must be carefully placed into the slot in the machine and the experiment can be run.
A singular peak in the resulting graph would mean that the particles are well dispersed in the medium and a zeta potential value, lower than -20 mV will indicate that there are stable carboxyl groups present in the solution. This negative charge is formed because the carboxyl groups will lose a proton and become negatively charged when present in aqueous solution
To obtain high-resolution images of our nanoparticles for size determination, we conducted Transmission Electron Microscopy (TEM) studies. TEM is a microscopy technique that works by transmitting a beam of electrons through a thin specimen and capturing the transmitted electrons to create high-resolution images. The sample must be loaded on a copper grid and air-dried to remove all the excess water. This sample can be stored in a microcentrifuge tube at room temperature if needed.
The binding of the probe to the carboxylated magnetic nanoparticle
Our probe structure was divided into three different single-stranded DNA oligos, that would bind together by complementarity, to form the working probe. These three sequences were named A, B and C. Out of these structures, B contains an NH2 group that could be bound to the carboxyl (COOH) group present in the nanoparticle by an amide bond formed by EDC coupling reaction.
This involves washing the nanoparticles with MES to prevent agglomeration, followed by treatment with EDC, which activates the carboxyl groups present in the nanoparticles. These activated carboxyl groups can now form an amide bond with sequence B. So, the required volume of sequence B is added to the activated carboxylated magnetic nanoparticles and incubated overnight at room temperature with constant shaking. After this step, the supernatant is removed and the required volume of sequences A and C are added and incubated for 1 hour. Now, The supernatant can be removed after pulling the particles down using a magnet and fresh medium can be added.
All the experiments mentioned below were performed on a Costar, black, flat-bottom plate.
The readout was taken by performing an emission scan, with a constant excitation of 490 nm and emission range of 500 to 600 nm, with readings taken in 1 nm steps.
We conducted this experiment to see if the binding was indeed giving us a fluorescence change. The results from this experiment could be used as a comparison to show the effect of binding the probes to the nanoparticle
This was performed to see if there was a considerable increase in fluorescence in the nanoprobe after miDNA was added. In the experiment, the amount of probe and total volume was kept constant. The control had an equal volume of the medium used for dilution of the miDNA as the volume of the miDNA sample.
This experiment was performed to see if our probes would be able to give a change in fluorescence intensity even at low levels of miDNA as the miRNA levels in the blood are in picomolar levels. We tested as low as 10 picomolar concentrations of miDNA. The volume of the probe and sample was kept constant and the concentration of the sample was varied.
We performed this experiment to see how specific our probe was as our probe should be able to differentiate between the target miRNA and other similar miRNA. For this, we ordered a miDNA with a difference in just two nucleotides, compared to our target miDNA. For the experiment, the amount of probe and total volume of solution was kept constant. We then made four samples for each probe, one with just media as the sample, one with a completely different miDNA as the sample, one with our target miDNA and one with the miDNA with just two nucleotide difference to the target miDNA.
All the experiments mentioned below were performed on a Greiner 96 Flat Bottom Transparent Polystyrene Cat. No.: 655101/655161/655192 [GRE96ft.pdfx] plate. The FluoroLog fluorometer is a versatile instrument that can be used to measure the fluorescence of a wide variety of samples. It is important to follow the protocol carefully to ensure accurate and reproducible results. If you are using a 200 microliter cuvette, you will need to use a cuvette adaptor. The cuvette adaptor allows the smaller cuvette to be used in the fluorometer.
This experiment was designed to find the ideal time for the FRET system to show maximum quenching. FRET works by transferring energy from a donor fluorophore to an acceptor quencher when the two molecules are close together. In this experiment, a fluorophore was attached to the aptamer and a quencher was attached to the cDNA. When the aptamer and cDNA bind together to form a duplex, the fluorophore and quencher are brought close together, and FRET occurs. The fluorescence intensity was measured over time to determine the ideal time for maximum quenching to occur.
In this experiment, the aptamer and cDNA were mixed together in specific proportions, such as 0.25 micromolar aptamer and 0.75 micromolar cDNA. The fluorescence intensity was then measured at the start of the experiment (time t=0) and every 10 minutes thereafter, until the fluorescence quenching did not change much over time.
This experiment was designed to determining the best ratio of aptamer to target molecule for the formation of an aptasensor.This experiment was performed on two different machines, a fluorometer and a microplate reader, to see if the results were similar. The goal was to determine if the trends observed were consistent across different machines, which would suggest that the results are reliable.
This experiment is a crucial test of the principle of the aptasensor, and it is designed to determine how sensitive the aptasensor is. The experiments were conducted on a microplate reader instead of a fluorometer because the researchers did not have enough reagents for the fluorometer. To conduct this experiment we select the most ideal incubation time and molar ratio for the formation the aptasensor and we take 150 microliter of the aptasensor and add 50 microliter of varying concentration of the biomarkers ranging from 10 ng/ml to 400 ng/ml mimicking the physiological range present in plasma.
Isothermal titration calorimetry (ITC) is a powerful technique for measuring the thermodynamics of biomolecular interactions, including the formation of DNA duplexes. It is a direct method that does not require any labeling or modification of the DNA strands. ITC is also very sensitive, allowing for the detection of even weak interactions.
There are several reasons why ITC is useful for checking duplex formation between two DNA strands:
ITC directly measures the enthalpy and binding affinity of the interaction. This information can be used to assess the stability of the DNA duplex and to understand the forces that drive duplex formation.
ITC does not require any labeling or modification of the DNA strands. This makes it a versatile technique that can be used to study a wide range of DNA duplexes, including those that are difficult or impossible to label.
ITC is very sensitive, allowing for the detection of even weak interactions. This is important for studying DNA duplexes, which can be relatively weak, especially for short strands.
The principle of isothermal titration calorimetry (ITC) is based on the direct measurement of the heat released or absorbed during a biomolecular binding event. ITC experiments are performed at a constant temperature, so the heat measured is directly proportional to the change in enthalpy of the binding reaction. ITC instruments typically consist of two cells: a sample cell and a reference cell. The sample cell contains the biomolecule of interest, while the reference cell contains a buffer solution. The two cells are enclosed in an adiabatic jacket, which prevents heat from being transferred to or from the environment.
During an ITC experiment, a ligand solution is titrated into the sample cell. When the ligand binds to the biomolecule, heat is either released or absorbed. This heat is detected by the calorimeter and recorded as a thermogram. The thermogram is a plot of the heat released or absorbed as a function of the amount of ligand added.
The thermogram can be analyzed to determine the following thermodynamic parameters of the binding reaction:
PCR is used to amplify a desired DNA sequence.These are temperature influenced enzymatic reactions.It consists mainly of 3 steps.
We ran a gradient PCR to find out the ideal annealing temperature of our primers. Gradient pcr allows to amplify multiples samples in different annealing temperature in a single run. By identifying the temperature that gives the highest yield of our desirable product we can find the optimal annealing temperature. It is similar to normal pcr amplification with respect to all other steps. The experiment was conducted in a Thermofischer SimpliAmp Thermal Cycler.
Ligation PCR is performed to confirm the ligation reaction..It helps ensure that the ligation ha been completed without errors like deletions, re-arrangements or insertions.It aslo helps to understand the orientation of inserted fragment relative to vector. Forward primer and reverse primers are designed specifically for both insert and vector; one being specific to the vector while the other being specific to the insert. Analyzing the bands obtained upon running agarose gel electrophoresis, aforementioned assumptions can be confirmed.
Colony pcr is a rapid and cost effective method for screening a large number of bacterial clones. Here, we performed it to confirm that the host cells have taken up the recombinant DNA and to identify the required clonal colony. Selecting suitable bacteria colonies is very crucial for further steps of cloning. Primer set specific to the insert is added such that only those sequences, if present in the plasmid gets amplified. The bands obtained on the agar gel is further analyzed to ensure that size of amplified DNA fragment is close to insert size and thus confirm that the transformation has occurred. We also included a positive control to verify the performance of pcr reaction and check for contamination.
Gel electrophoresis helps to separate the samples according to size in agarose gel.The samples are pipetted in to the wells of the gel after mixing with a loading dye (necessary for visualization). An electric current is applied to the Anode region causing the negatively charged dna to move towards the Cathode region.
A molecular weight ladder is also loaded into a well along with the samples while running the gel. Obtained bands are compared against the bands of the ladder to determine size. The size is proportional to migration rate.Smaller fragments move quickly toward the bottom region.
Ethidium Bromide is used to visualize the DNA in gel. The DNA sample takes up the dye as they migrate through the gel. Pale pink bands are observed on illumination with UV light once the electrophoresis is completed.
We mainly performed Gel electrophoresis as a confirmation step for numerous experiments like PCR amplification reactions, restriction digestions, ligation reactions and transformations. All electrophoresis experiments were run on the Bio - Rad Horizontal Electrophoresis System.
During restriction digestion, specific restriction enzymes - usually endonucleases- cut the dna fragments at specific locations called recognition sites.
In order to introduce our amplified insert into the vector pET19b, we need to create sticky ends on both insert and vector by restriction-digesting both DNA sequences using the same restriction enzymes. It is very crucial to select appropriate restriction enzymes for cutting both vector and insert. We used two different enzymes, one for each cutting each in order to decrease chances of non-specific cutting and ensure proper orientation of the insert in the vector. Restriction digestion was performed using BamH1 and Nde1, both high fidelity endonucleases.High fidelity(HF) enzymes have a higher accuracy rate.Appropriate incubation time was set, since excess incubation can lead to over digestion and formation of excessive DNA fragments.
Ligation reaction is performed to join the compatible ends of the insert and vector after restriction digestion. After successful ligation, we obtain the recombinant DNA molecule - a circular plasmid- that can be transformed into host cells.
We performed the reaction using T4 Ligase, a versatile ligase that ligates DNA fragments by catalyzing the formation of phosphodiester bonds between juxtaposed 5' phosphate and 3' hydroxyl termini in double-stranded DNA using ATP as a coenzyme.
During bacterial transformation, bacteria takes up a naked DNA (foreign gene) from its environment.This is a horizontal gene transfer process. Not all bacteria can take up naked DNA. Bacteria is made artificially competent using chemical or electrical pulses.Transformation experiments are generally performed in a Laminar Air Flow(LAF) or Biosafety Cabinets with sterile equipments and reagents to reduce the risk of contamination.
We are required to perform two transformations.
The first transformation is performed to introduce the recombinant DNA molecule into Dh5α strain of E.coli create in order to create multiple copies of the rDNA. Dh5α is known for its high tansformation efficiency. Since the natural competency of E. coli is very low or even nonexistent, the cells are made competent for transformation by heat shock. We also included a negative control for assessing the efficiency and specificity of transformation.
The second transformation is performed in order to transform the isolated plasmid into BL21 strain of E.Coli cells. BL21 is an E.coli strain commonly used for expression of recombinant protein due to its modified T7 RNA Polymerase system, high expression levels owing to high affinity of T7 RNA polymerase for it’s promoter and comparatively fast growth rate.
Alkaline lysis is used to separate plasmid DNA from other cell constituents,including genomic DNA. This is done to isolate the transformed pET19b plasmid which is then transformed into BL21 strain of E.coli for expression of protein.
Alkali like 10N Naoh is added to the resuspended cells for cell lysing; a detergent like SDS is used to denature most of the cell protein. Cell debris is precipitated using potassium acetate which also helps in renaturing the double stranded plasmid DNA. This is then dissolved which is eluted after centrifuging. Final plasmid DNA precipitation is then carried out using ethanol/isopropanol.