The DNA extraction process for Colletotrichum falcatum in the Point-of-Care (POC) red rot detection system is designed to be efficient and suitable for field diagnostics. It begins with the preparation of a lysis buffer containing 0.5 M NaCl, 1% SDS, 10 mM Tris-HCl, and 1 mM EDTA, which helps break down the tough fungal cell walls and releases the genomic material. Before use, the buffer is heated to 65°C to ensure the SDS dissolves properly, optimizing its lytic activity.

Once the fungal cells are lysed, the solution is treated with a phenol-chloroform-isoamyl alcohol (PCI) mixture in a 25:24:1 ratio. This step is crucial for protein denaturation and separation of cellular debris from the DNA. The PCI mix is carefully prepared with Tris-HCl and stored in a dark, tightly capped bottle at 4°C to maintain its efficacy. This organic extraction step ensures the removal of proteins and lipids, leaving purified nucleic acids in the aqueous phase.

Following PCI treatment, the sample is further purified using a chloroform-isoamyl alcohol (24:1) solution, which removes any remaining phenol and enhances DNA purity. This solution is also stored in a dark bottle at 4°C to prevent degradation.

The final step involves resuspending the extracted DNA in T0.1E buffer (10 mM Tris-HCl, 0.1 mM EDTA) to stabilize and store the DNA. This buffer protects the DNA from degradation while ensuring it remains suitable for downstream applications like amplification via LAMP, which is essential for sensitive detection of the pathogen in early stages.

This method balances cost-effectiveness, reliability, and simplicity, making it an ideal choice for developing a rapid and field-ready diagnostic tool for red rot detection in sugarcane.

Initial Equilibration: The experiment begins with the ligand (typically in the syringe) being titrated into the sample cell containing the macromolecule (usually a protein or nucleic acid). Both cells contain the same buffer to maintain constant conditions.**

Titration: Small volumes of the ligand solution are injected into the sample cell in a controlled manner, typically in a series of steps. As the ligand mixes with the macromolecule, a binding reaction takes place.

Heat Exchange Measurement: Heat is either absorbed or released during the binding reaction, leading to temperature changes in the sample cell. The calorimeter detects these temperature changes and converts them into heat flow data.

Data Analysis: The heat flow data are then analyzed to determine the binding parameters, including the binding constant (Kd), enthalpy change (ΔH), entropy change (ΔS), and stoichiometry (n).

Determination of Binding Affinities: ITC is widely used to measure the binding affinities between various biomolecules, such as proteins, nucleic acids, and small molecules. It can help identify the strength of molecular interactions, which is crucial in drug discovery and understanding biological processes

Enzyme Kinetics: ITC can be used to study enzyme-substrate interactions and determine kinetic parameters, such as the Michaelis-Menten constant ($K_m$) and turnover number ($K_{cat}$).

Thermodynamic Studies: ITC provides insights into the thermodynamics of binding reactions, including ΔH and ΔS, which can shed light on the forces driving molecular interactions.

Stoichiometry Determination: ITC can determine the stoichiometry of complex formation, i.e., the number of ligand molecules binding to a macromolecule. Screening for Ligand Binding: ITC can be used to screen potential drug candidates for their binding affinity to a target protein, helping in drug development.

In summary, isothermal titration calorimetry is a versatile technique that allows researchers to quantitatively study molecular interactions by measuring heat changes during a binding reaction. It is invaluable in understanding the thermodynamics of these interactions, which has wide-ranging applications in biology, chemistry, and pharmaceutical research.

Phage display is a technique used to express specific proteins by utilizing a bacteriophage (a virus that infects bacteria) as a host. This method finds applications in various fields, including virology, structural biology, and molecular biology, particularly when the goal is to display proteins on the surface of the phage. By inserting the gene for the ZAP1 protein into the pIII coat protein gene region, the desired protein can be expressed on the surface of the virion. Here’s a detailed overview of how the phage display operates.

Genetic Modification: The gene encoding a foreign protein is fused to the gene of a phage coat protein, typically the pIII protein. This fusion allows the foreign protein to be displayed on the surface of the phage.

Phage DNA Preparation: The modified phage DNA is constructed by inserting the foreign gene into the phage genome using transformation. This results in a recombinant phage DNA that encodes both the phage coat proteins and the foreign protein.

Transfection: The modified phage DNA is introduced into bacterial cells (most commonly E. coli) through a process called transfection. During this step, the bacterial cells take up the modified DNA, leading to the expression of the fusion protein.

Phage Production: Once inside the bacteria, the phage genes are expressed, and new phage particles are assembled. The foreign protein is displayed on the surface of these newly formed phage particles, which are then released into the culture medium.

Amplification: The bound phages are collected and used to infect fresh bacterial cells. This step amplifies the population of phages that display the desired foreign protein. As the bacteria replicate, they produce a new batch of phages, again displaying the selected proteins.