Experiments

The 14 Minute Transformation Protocol for NEB 5-alpha and 10-beta competent E. coli offers a balance between speed and efficiency, making it suitable for routine cloning with higher transformation efficiency than ultra-fast protocols. The protocol includes critical steps such as ice incubation, heat shock, and outgrowth to maximize colony formation. The following protocol results in only 25% efficiency compared to the High Efficiency Transformation Protocol. For C2987H, perform steps 1-6 in the tube provided.

1. Prepare all required reagents and consumables:
   • NEB 5-alpha or 10-beta competent cells (thawed on ice)
   • Plasmid DNA (1 pg–100 ng in 1–5 μl)
   • SOC or NEB 10-beta/Stable Outgrowth Medium (950 μl, room temperature)
   • Sterile 1.5 ml microtubes
   • Ice and ice bucket
   • Water bath at 42°C
   • 37°C incubator
   • Sterile pipettes and tips
   • LB agar plates with appropriate antibiotic
2. Procedure (C2987H/C2987I)
   • For C2987H: Remove cells from -80°C freezer and thaw in your hand. For C2987I: Thaw a tube of NEB 5-alpha Competent E. coli cells on ice until the last ice crystals disappear. Mix gently and carefully pipette 50 μl of cells into a transformation tube on ice.
   • Add 1-5 μl containing 1 pg-100 ng of plasmid DNA to the cell mixture. Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.
   • Place the mixture on ice for 10 minutes. Do not mix.
   • Heat shock at exactly 42°C for exactly 30 seconds. Do not mix.
   • Place on ice for 3 minutes. Do not mix.
   • Pipette 200 μl of room temperature SOC into the mixture. Immediately spread 50-100 μl onto a selection plate and incubate overnight at 37-42°C. NOTE: Selection using antibiotics other than ampicillin may require some outgrowth before plating on selective media. Colonies develop faster at temperatures above 37°C, however some constructs may be unstable at elevated temperatures.

The High Efficiency Transformation Protocol by NEB optimizes the uptake of plasmid DNA into E. coli strains 5-alpha and 10-beta, achieving efficiencies of 1–3 × 109 cfu/μg. It involves heat shock (42°C, 30 seconds) and recovery in specialized media, ideal for cloning large plasmids or BACs (10-beta).

1. Prepare all required reagents and consumables:
   • NEB 5-alpha or 10-beta competent cells (thawed on ice)
   • Plasmid DNA (1 pg–100 ng in 1–5 μl)
   • SOC or NEB 10-beta/Stable Outgrowth Medium (950 μl, room temperature)
   • Sterile 1.5 ml microtubes
   • Ice and ice bucket
   • Water bath at 42°C
   • 37°C incubator
   • Sterile pipettes and tips
   • LB agar plates with appropriate antibiotic
2. Procedure (C2987H/C2987I, for C2987H, perform steps 1-7 in the tube provided)
   • For C2987H: Thaw a tube of NEB 5-alpha Competent E. coli cells on ice for 10 minutes. For C2987I: Thaw a tube of NEB 5-alpha Competent E. coli cells on ice until the last ice crystals disappear. Mix gently and carefully pipette 50 μl of cells into a transformation tube on ice.
   • Add 1-5 μl containing 1 pg-100 ng of plasmid DNA to the cell mixture. Carefully flick the tube 4-5 times to mix cells and DNA. Do not vortex.
   • Place the mixture on ice for 30 minutes. Do not mix.
   • Heat shock at exactly 42°C for exactly 30 seconds. Do not mix.
   • Place on ice for 5 minutes. Do not mix.
   • Pipette 950 μl of room temperature SOC into the mixture.
   • Place at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
   • Warm selection plates to 37°C.
   • Mix the cells thoroughly by flicking the tube and inverting, then perform several 10-fold serial dilutions in SOC.
   • Spread 50-100 μl of each dilution onto a selection plate and incubate overnight at 37°C. Alternatively, incubate at 30°C for 24-36 hours or 25°C for 48 hours.

Colony PCR is a rapid screening method used to verify whether bacterial colonies contain the expected plasmid insert. Instead of isolating plasmid DNA, bacterial cells from individual colonies are lysed directly and used as template in a PCR reaction. This approach saves time and allows quick identification of positive clones. It is widely used after transformations, though positive results should always be confirmed by sequencing.

1. Prepare all required reagents and consumables:
   • Agar plates with selective antibiotic (colonies to be tested)
   • Sterile pipette tips and pipettes
   • Microcentrifuge tubes (1.5mL) and PCR tubes
   • Sterile nuclease-free water
   • PCR Master Mix (commercial 2× mix or individual components: polymerase, buffer, dNTPs)
   • DNA polymerase (RUN, A&A)
   • Buffer for PCR (RUN, A&A)
   • dNTPs
   • Forward and reverse primers specific to the insert/construct
   • Thermocycler
   • Agarose gel electrophoresis setup (gel, DNA ladder, staining solution, power supply, UV or blue-light imager)
   • Laboratory safety equipment: gloves, lab coat, goggles
2. Prepare primers and reagents
   • Design primers to amplify the region of interest (insert-specific or vector flanking primers).
   • Prepare the PCR Master Mix according to manufacturer’s instructions.
3. Pick bacterial colonies
   • Touch a single colony with a sterile pipette tip.
   • Resuspend the cells in a small volume of sterile water (100μL) or buffer in a microcentrifuge tube.
4. Lyse the cells
   • Perform cell lysis by heat treatment: 10 min in 95°C
   • Use the resulting lysate as template DNA
5. Set up PCR reactions
   • Aliquot all of the ingredients of PCR into PCR tubes. Use the table below to calculate the volumes.

   Reagent	Volume
   DNA template	2 μL
   Buffer	2 μL
   dNTPs	0.4 μL
   Primer Forward	1 μL
   Primer Reverse	1 μL
   DNA polymerase	0.2 μL
   Distilled water	13.4 μL

   • Include a negative control (no template) and a positive control (known plasmid).
6. Run PCR
   • Place the reactions in a thermocycler. Set the program with settings listed below:

   Step	Temparature	Time	Cycles
   Initial denaturation	95°C	3 min	1
   Denaturation	95°C	30 s	35
   Annealing	55°C	30 s	35
   Elongation	72°C	60 s	35
   Final extension	72°C	5 min	1
   Hold	4°C	∞	1

   • Run the cycling program optimized for your polymerase and primers (denaturation, annealing, extension).
7. Analyze products
   • Load PCR products on an agarose gel with a DNA ladder.
   • Compare observed band sizes with the expected amplicon.
   • Positive clones can be confirmed by sequencing.

This protocol describes how to prepare chemically competent E. coli cells capable of efficiently taking up plasmid DNA through heat shock transformation. Using calcium chloride (CaCl2) treatment, the cells become permeable to DNA, achieving transformation efficiencies typically ranging from 106 to 108 colony-forming units (cfu) per microgram of DNA, depending on the strain and conditions.

1. Prepare all required reagents and consumables:
   • E. coli strain (e.g., DH5α, BL21)
   • LB or SOB broth (sterile)
   • Ice-cold 0.1M Calcium chloride (CaCl2) solution (sterile)
   • Ice-cold 15% glycerol solution (sterile)
   • Shaking incubator at 37°C
   • Refrigerated centrifuge (4°C)
   • Ice bucket
   • Sterile 50mL and 1.5mL centrifuge tubes
   • Sterile pipettes and tips
2. Starter Culture: Inoculate a single colony of E. coli into 2–5mL LB broth. Incubate overnight at 37°C with shaking (200–250 rpm).
3. Main Culture: Dilute 1mL of the overnight culture into 100mL fresh LB or SOB medium. Grow at 37°C with shaking until the culture reaches an OD600 of approximately 0.4–0.6 (about 2–3 hours).
4. Cooling: Place the culture on ice for 15–30 minutes to chill the cells.
5. Centrifugation: Centrifuge the culture at 2500–4000 × g for 10 minutes at 4°C. Carefully discard the supernatant.
6. Calcium Chloride Wash: Gently resuspend the cell pellet in 25–50mL of ice-cold 0.1M CaCl2 solution. Incubate on ice for 30 minutes. Centrifuge again under the same conditions and discard the supernatant.
7. Final Resuspension: Resuspend the pellet in 2–5mL of ice-cold 0.1M CaCl2 containing 15% glycerol. Mix gently to avoid shearing the cells.
8. Aliquoting and Storage: Dispense the competent cells into sterile microcentrifuge tubes (50– 200μL per tube). Immediately flash-freeze the aliquots in a -80°C freezer.

Introduction

This protocol provides an optimized and standardized approach for the maintenance of HEK293T, WI-38, HepG2, and A549 cell lines, ensuring optimal viability and reproducibility. It covers cell thawing, passaging, transfection and cryopreservation, with tailored guidelines that incorporate specific modifications to accommodate the unique growth characteristics and requirements of each cell type.

Safety note

This protocol provides an optimized and standardized approach for the maintenance of HEK293T, WI-38, HepG2, and A549 cell lines, ensuring optimal viability and reproducibility. It covers cell thawing, passaging, transfection and cryopreservation, with tailored guidelines that incorporate specific modifications to accommodate the unique growth characteristics and requirements of each cell type.

Characteristics of cell lines

HepG2

HepG2 is a human hepatocellular carcinoma cell line derived from the liver tissue of a 15-year-old male with hepatoblastoma. These cells are widely used as an in vitro model of human hepatocytes due to their ability to retain many liver-specific metabolic and synthetic functions, including the expression of plasma proteins and certain cytochrome P450 enzymes. Being a tumor-derived cell line, HepG2 cells are immortal and proliferate indefinitely under standard culture conditions.

HEK293T

HEK293T cells are a derivative of the human embryonic kidney 293 (HEK293T) cell line, originally established from embryonic kidney tissue transformed with adenovirus 5 DNA. The “T” variant is stably transfected with the SV40 large T antigen, enabling episomal replication of plasmids containing the SV40 origin of replication. HEK293T cells are immortalized and tumorigenic, and they are extensively employed in molecular biology and biotechnology, particularly for high-efficiency transient transfection and recombinant protein or viral particle production.

A549

A549 is a human lung carcinoma cell line derived from the alveolar basal epithelial cells of a 58-year-old Caucasian male with adenocarcinoma. These cells exhibit epithelial morphology and retain many features of alveolar type II pneumocytes, including the ability to synthesize and secrete pulmonary surfactant components. A549 cells are cancer-derived and therefore immortal, and they are frequently used as a model for respiratory epithelium, lung cancer biology, and drug testing.

WI-38

WI-38 is a normal human diploid fibroblast cell line derived from lung tissue of a 3-month-old female fetus. Unlike the previously mentioned tumor-derived lines, WI-38 cells are non-tumorigenic and exhibit a finite lifespan in culture, undergoing replicative senescence after approximately 50 population doublings (the Hayflick limit). WI-38 has been widely used in research on cellular aging, vaccine production, and normal human cell biology, serving as a reference for comparison with immortalized or transformed cell lines.

Thawing cells

Goal of the experiment

The goal of this experiment is to efficiently and aseptically thaw cryopreserved cells, restoring their viability and functionality for subsequent culture. This procedure ensures minimal cellular stress and maximizes post-thaw recovery by rapidly transitioning cells from cryogenic storage to optimal growth conditions.

1. Prepare all required reagents and consumables:
   • Complete culture medium - DMEM + 10% FBS + 1× P/S + 1× GlutaMAX
   • Cryovial containing frozen cells of appropriate cell line
   • Tissue culture dish Ø 60 mm
   • Pipet controller and sterile serological pipettes (5 mL and 10 mL)
   • Pipet and sterile 1 mL tips
   • 15 mL centrifuge tube
   • Centrifuge compatible with 15 mL tubes
   • Brightfield (inverted) microscope
   • 37°C water bath
   • CO2 incubator set to 37°C and 5% CO2
2. Procedure
   
   Important: During thawing cells are in a vulnerable state. Before you start prepare all the materials, set the pipet for 1 mL, label tissue culture dish, prepare 5 mL of complete culture medium in a centrifuge tube and keep the frozen cryovial on dry ice.
   
   
   • Preheat complete culture medium in warm water bath to 37°C
   • Put the cryovial containing cells in a warm water bath at 37°C, making sure that the lids stay out of the water. It should take no more than 2 min.
   • When only a small piece of frozen cell suspension remains in the cryovial, immediately transfer it under sterile conditions to the biosafety cabinet and gently pipette the contents into a pre-warmed 15 mL centrifuge tube containing preheated 5 mL of complete culture medium.
   • Immediately spin the cells down in the centrifuge at appropriate settings.

   Cell line	centrifuge settings
   HepG2	100 × g, 8 min, RT
   HEK293T	500 × g, 5 min, RT
   A549	500 × g, 5 min, RT
   WI-38	100 × g, 8 min, RT

   • Carefully discard the medium. Resuspend the cells in 4 mL of medium.
   • Transfer the suspended cells onto Ø 60 mm tissue culture dish.
   • Incubate cells in standard conditions: 37°C, 5% CO2.
   • Cells should be passed when they reach approximately 80% confluency (about 1 day) onto a Ø 100 mm culture dish. For experimental use, cells are recommended to be utilized after minimum two passages to ensure phenotypic stability and optimal viability.

Passing cells

Goal of the experiment

The aim of this experiment is to subculture adherent cell lines under optimal and aseptic conditions to maintain their viability, phenotypic stability, and experimental reproducibility. Proper passaging prevents overconfluence, senescence, and phenotypic drift, ensuring consistent cellular behavior across experiments.

1. Prepare all required reagents and consumables:
   • Complete culture medium - DMEM + 10% FBS + 1× P/S + 1× GlutaMAX
   • Trypsin-EDTA
   • PBS
   • Cell culture with confluence of about 80%
   • Tissue culture dish Ø 100 mm
   • Pipet controller and sterile serological pipettes (5 mL and 10 mL)
   • Pipet and sterile 1 mL tips
   • Brightfield (inverted) microscope
   • 37°C water bath
   • CO2 incubator set to 37°C and 5% CO2
2. Procedure
   
   
   • Preheat complete culture medium in warm water bath to 37°C
   • Check confluency of cells using the brightfield microscope. Cells should cover about 80% of the tissue culture dish.
   • Carefully pipette out the old growth medium from the corner of the dish.
   • Gently wash the cells with an appropriate amount of PBS by pouring it onto the walls. Swirl it around and pipette it out from the corner of the dish.
   • Add the appropriate amount of Trypsin-EDTA, it should cover the surface. Incubate for a maximum of 10 min in standard conditions. Check dissociation every 2 mins under the brightfield microscope.
   • After all cells dissociate, add a complete culture medium to neutralize the Trypsin-EDTA. Suspend the cells.
   • Optional: Prior to plating for the experiment, cells should be centrifuged to eliminate residual trypsin-containing medium.
   • Determine the amount or ratio of cells that should be added to the new dish. For standard passing every 2 or 3 days ratio is listed below.

   Cell line	centrifuge settings	ratio
   HepG2	100 × g, 8 min, RT	1:3 – 1:4
   HEK293T	500 × g, 5 min, RT	1:4 – 1:6
   A549	500 × g, 5 min, RT	1:4 – 1:5
   WI-38	100 × g, 8 min, RT	1:2 – 1:3

   Reagent	100 mm	60 mm	6-well	24-well	96-well
   Volume of medium in which the culture is carried out [mL]	10	4	2	0.4	0.1
   Volume of PBS used to wash the cells [mL]	11	5	3	0.5	0.15
   The volume of Trypsin-EDTA used to detach the cells [mL]	1.5	0.5	0.25	100	0.05

Cryopreservation

Goal of the experiment

The goal of this experiment is to cryopreserve viable and functionally stable cell stocks of HEK293T, THLE-2, HepatoXcell, and HepG2 lines for long-term storage. By freezing cells at optimal confluency in appropriate cryoprotective media under controlled-rate cooling conditions, this protocol ensures minimal cellular damage and maximizes post-thaw recovery and reproducibility in future experiments.

1. Prepare all required reagents and consumables:
   • Complete culture medium - DMEM + 10% FBS + 1× P/S + 1× GlutaMAX
   • Freezing medium - 90% complete DMEM + 10% DMSO
   • Trypsin-EDTA
   • PBS
   • Cryovials
   • Freezing container
   • Cell culture with confluence of about 80%
   • Pipet controller and sterile serological pipettes (5 mL and 10 mL)
   • Pipet and sterile 1 mL tips
   • 15 mL centrifuge tube
   • Brightfield (inverted) microscope
   • 37°C water bath
   • CO2 incubator set to 37°C and 5% CO2
2. Procedure
   
   Important: During cryopreservation, after suspending cells in freezing medium, cells are in a vulnerable state. Before you start prepare all the materials, set the pipet for 1 mL, label cryovials, prepare freezing medium and freezing container. Work fast and immediately after loading the freezing container transfer it into the -80°C freezer.
   
   
   • Preheat complete culture medium in a warm water bath to 37°C, prepare appropriate amount and type of freezing medium (90% complete DMEM + 10% DMSO).
   • Check confluency of cells using the brightfield microscope. Cells should cover about 80% of the tissue culture dish.
   • Carefully pipette out the old growth medium from the corner of the dish.
   • Gently wash the cells with an appropriate amount of PBS by pouring it onto the walls. Swirl it around and pipette it out from the corner of the dish.
   • Add the appropriate amount of Trypsin-EDTA, it should cover the surface. Incubate for a maximum of 10 min in standard conditions. Check dissociation every 2 mins under the brightfield microscope.
   • After all cells dissociate, add a complete culture medium to neutralize the Trypsin-EDTA. Suspend the cells.
   • Transfer cells to the centrifuge tube and spin the cells down in the centrifuge at appropriate settings. Discard the supernatant.
   • Resuspend the cells in freezing medium and immediately distribute 1 mL of the suspension into each cryovial.

   Cell line	centrifuge settings
   HepG2	100 × g, 8 min, RT
   HEK293T	500 × g, 5 min, RT
   A549	500 × g, 5 min, RT
   WI-38	100 × g, 8 min, RT

Transfection

Introduction

LipofectamineTM 3000 (Thermo Fisher) is a lipid-based transfection reagent optimized for efficient delivery of plasmid DNA into mammalian cells, including hard-to-transfect lines. Together with the P3000TM enhancer, it promotes complex formation between DNA and cationic lipids, resulting in high transfection efficiency and low cytotoxicity. Complexes are prepared in serum-free medium (Opti-MEMTM) and directly added to adherent cells in culture without the need to replace the medium after transfection. This protocol describes how to prepare DNA–lipid complexes and transfect adherent cells in 24-well plates. Conditions can be scaled up or down depending on the culture vessel.

1. Prepare all required reagents and consumables:
   • Cells and culture
     • Mammalian cells for transfection
     • Complete growth medium appropriate for the chosen cell line
     • 24-well tissue culture plates (or other format)
     • PBS (without Ca2+/Mg2+)
     • Trypsin or dissociation reagent
   • Transfection reagents
     • LipofectamineTM 3000 (Thermo Fisher)
     • P3000TM reagent (Thermo Fisher)
     • Opti-MEMTM Reduced Serum Medium
     • Plasmid DNA to be transfected (purified, endotoxin-free)
     • Plasmid DNA to be transfected (purified, endotoxin-free)
   • • Sterile 1.5 mL microcentrifuge tubes
     • Filtered sterile pipette tips (P10, P200, P1000)
     • Micropipettes (P10, P200, P1000)
     • Biological safety cabinet (laminar flow hood)
     • CO2 incubator (37°C, humidified atmosphere)
     • Vortex mixer and benchtop centrifuge
     • Inverted microscope for cell morphology check
     • Personal protective equipment (lab coat, gloves, goggles)
2. Procedure
   
   Note: Seed cells one day prior to transfection so they reach ~70–90 % confluency on the day of transfection.
   
   
   • Prepare DNA solution and Lipofectamine 3000 solution
     • Use the amounts listed in the table below to prepare both solutions.
     • Mix gently by pipetting.

   	Solution 1			Solution 2
   Reagent	Opti-MEM	P3000TM	DNA	Opti-MEM	LipofectamineTM 3000
   Volume per 1 well (24-well plate)	25 uL	1 uL	500 ng	25 uL	1,5 uL

   Form DNA-lipid complexes
   • Combine the DNA/P3000TM solution with the LipofectamineTM 3000 solution.
   • Mix gently by pipetting up and down.
   • Incubate at room temperature for 10–15 min to allow complex formation.
   Add complexes to cells
   • Add the entire portion of complexes dropwise to each well containing cells in complete growth medium.
   • Gently rock the plate to evenly distribute the complexes.
   Incubation
   • Return the cells to the CO2 incubator (37°C, humidified, 5 % CO2).
   • Do not change the medium; complexes remain with the cells throughout the incubation.
   Post-transfection analysis
   • After 24–48 h, assess transfection efficiency (e.g. via reporter gene expression or downstream assay).
   • Monitor cell morphology and viability using a microscope or flow cytometry.

This protocol describes the preparation of adherent HepG2 (hepatocellular carcinoma) and HEK (human embryonic kidney) cells cultured in 24-well plates for flow cytometry analysis.

1. Prepare all required reagents and consumables:
   • Adherent HepG2 or HEK cell cultures in 24-well plates (70-90% confluent)
   • Phosphate-buffered saline (PBS), calcium and magnesium-free, sterile
   • 0.25% trypsin-EDTA solution (pre-warmed to 37°C)
   • Complete growth medium containing serum (pre-warmed to 37°C)
   • PBS buffer (cold, 4°C)
   • PBS buffer (room temperature)
   • 15 mL conical centrifuge tubes
   • Hemocytometer or automated cell counter
2. Procedure (C2987H/C2987I)
   • Pre-Protocol preparation
     • Pre-warm 0.25% trypsin-EDTA solution and complete growth medium to 37°C
     • Keep PBS buffer cold at 4°C
     • Label conical centrifuge tubes for each sample
     • Ensure cells are 70-90% confluent before harvesting
   • Cell washing
     • Remove culture medium from each well by gentle aspiration
     • Rinse wells with 500 μL sterile, room temperature PBS to remove residual serum
   • Enzymatic Detachment
     • Add 200 μL of pre-warmed 0.25% trypsin-EDTA solution per well
     • Incubate plates at 37°C for 5 - 8 minutes
       • Monitor cell detachment under inverted microscope
       • Gently tap or rock the plate if needed to facilitate detachment
   • Trypsin Neutralization and Cell Collection
     • Add 500-800 μL complete growth medium containing serum to each well to neutralize trypsin
     • Pipette cell suspension up and down 3-5 times to ensure single-cell suspension
     • Transfer cell suspension from each well into labeled 15 mL conical centrifuge tubes
   • Cell Washing and Preparation
     • Centrifuge tubes at 500 × g for 5 minutes at room temperature
     • Carefully discard supernatant without disturbing cell pellet
     • Resuspend pellet in 2 mL cold PBS buffer
     • Centrifuge again under the same conditions
     • Discard supernatant and resuspend pellet in 1 mL cold PBS buffer
   • Cell Count
     • Perform cell count using hemocytometer with trypan blue
     • Assess cell viability (should be >90% for optimal results)
   • Final Preparation for Flow Cytometry
     • Keep cells on ice (4°C) until analysis
     • Use cells for flow cytometry within 2-4 hours of preparation for best results

Based on Golden Gate Assembly Protocol for BsaI-HF v2 (NEB R3733)

1. Prepare all required reagents and consumables:
   • T4 DNA Ligase (NEB #M0202)
   • BsaI-HFv2 (NEB #R3733)
   • plasmid with backbone
   • NEB 10-beta competent E. coli
2. Procedure
   • Set up 25μL assembly reactions as follows:

   Reagent	Assembly reagents
   backbone vector	75 ng
   1-3 inserts	2:1 molar ratio to backbone
   T4 DNA Ligase Buffer (10X)	2.5 μl
   T4 DNA Ligase (NEB #M0202), 2000 U/μl	0.5 μl (1000 units)
   T4 DNA Ligase (NEB #M0202), 2000 U/μl	1.5 μl (30 units)
   Nuclease-free H2O	to 25 μl

   • Mix gently by pipetting up and down 4 times.
   • Briefly microcentrifuge (1 sec.) to bring material to the bottom of tube.
   • Transfer to thermocycler and program as follows: (5 min 37°C → 5 min 16°C) x 30 cycles followed by 5 min 60°C. If reactions are done overnight, add a 4°C terminal hold to the protocol, but repeat the final 5 min 60°C step the next day before the transformations.

Based on Golden Gate Assembly Protocol for SapI (NEB #R0569)

1. Prepare all required reagents and consumables:
   • T4 DNA Ligase (NEB #M0202)
   • SapI (NEB #R0569)
   • Destination Plasmid
   • NEB 10-beta Competent E. coli
2. Procedure
   • Set up 20μL assembly reaction as follows:

   Reagent	Assembly reagents
   Destination vector	3 nM (final concentration)
   Amplicon inserts	3 nM each amplicon (final concentration)
   SapI (NEB #R0569), 10 U/μl	1.5 μl (15 units)
   T4 DNA Ligase (NEB #M0202), 2000 U/μl	0.25 μl (500 units)
   T4 DNA Ligase Buffer (NEB #B0202) (10X)	2 μl
   Nuclease-free H2O (NEB #B1500)	to 20 μl

   • (37°C, 5 min → 16°C, 5 min) x 30 → 60°C, 5 min → 4°C.
   • Cool reaction to 4°C prior to transformation, or store completed assembly reactions at -20°C.

T4 DNA ligase is a crucial ATP-dependent enzyme utilized in molecular biology for joining DNA fragments. This enzyme was isolated from bacteriophage T4 and catalyzes the formation of phosphodiester bonds between adjacent 3'-OH and 5'-phosphate termini in double-stranded DNA.

1. Prepare all required reagents and consumables:
   • NEB T4 DNA Ligase
     • T4 DNA Ligase
     • 10X T4 DNA Ligase Reaction Buffer
   • Linearized DNA vector
   • DNA insert
   • RNase free water
2. Procedure
   • Set up the following reaction in a microcentrifuge tube on ice. Note: T4 DNA Ligase should be added last. Note that the table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes.

   Reagent	Assembly reagents	Comments
   Component	20 μl reaction
   T4 DNA Ligase Buffer (10X)*	2 μl	* The T4 DNA Ligase Buffer should be thawed and resuspended at room temperature.
   Vector DNA (4 kb)	50 ng (0.020 pmol)
   Insert DNA (1 kb)	37.5 ng (0.060 pmol)
   Nuclease-free water	to 20 μl
   T4 DNA Ligase	1 μl

   • Gently mix the reaction by pipetting up and down and microfuge briefly.
   • For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes.
   • For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours (alternatively, high concentration T4 DNA Ligase can be used in a 10 minute ligation)
   • Heat inactivate at 65°C for 10 minutes.
   • Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells.

HiFi assembly protocol is designed for multiple fragments seamless assembly. Based on: NEB HiFi DNA Assembly reaction protocol.

1. Prepare all required reagents and consumables:
   • Proper vector
   • Proper inserts
   • NEBuilder HiFi DNA Assembly Cloning Kit
2. Procedure
   • Set up reaction on ice:

   Reagent	Assembly reagents	Comments
   Recommended DNA Molar Ratio	vector:insert = 1:2
   Total Amount of Fragments	X μl (0.03–0.2 pmols)	Optimized cloning efficiency is 50–100 ng of vector with 2-fold excess of each insert. Use 5-fold molar excess of any insert(s) less than 200 bp. Total volume of unpurified PCR fragments in the assembly reaction should not exceed 20%. To achieve optimal assembly efficiency, design 15-20 bp overlap regions between each fragment.
   NEBuilder HiFi DNA Assembly Master Mix	10 μl
   Nuclease-free Water	10-X μl
   Total Volume	20 μl

   • Incubate samples in a thermocycler at 50°C for 15 minutes.
   • Following incubation, store samples on ice or at –20°C for subsequent transformation.
   • Transform to proper bacterial strains (e.g. NEB 10-beta competent E. coli)

Digestion for insert verification

1. Prepare all required reagents and consumables:
   • Plasmid DNA (500ng–1μg per reaction)
   • BsaI-HF®v2 (NEB R3733)
   • 10X rCutSmartTM Buffer
   • mQ water
2. Procedure
   • Set up reaction on ice:

   Components	50 μl reaction
   DNA	1 μg
   10X rCutSmartTM Buffer	5 μl
   BsaI-HF®v2 (NEB R3733)	1 μl
   Nuclease-free Water	to 50 μl

   • Gently mix the reaction by pipetting up and down and microfuge briefly.
   • Incubate at 37°C for 1 hour.
   • Denaturate for 20 minutes at 65°C

Digestion for insert verification

1. Prepare all required reagents and consumables:
   • Plasmid DNA (500ng–1μg per reaction)
   • BsaI-HF®v2 (NEB R3733)
   • 10X rCutSmartTM Buffer
   • mQ water
2. Procedure
   • Set up reaction on ice:

   Components	50 μl reaction
   DNA	1 μg
   10X rCutSmartTM Buffer	5 μl
   SapI	1 μl
   Nuclease-free Water	to 50 μl

   • Gently mix the reaction by pipetting up and down and microfuge briefly.
   • Incubate at 37°C for 1 hour.
   • Denaturate for 20 minutes at 65°C

The HiScribe T7 High Yield RNA Synthesis Kit (NEB) is designed for efficient in vitro transcription (IVT) of RNA from a DNA template containing a T7 promoter. This kit enables the synthesis of high yields of RNA in a short reaction time, and is commonly used in applications such as mRNA production, probe synthesis, or synthetic biology constructs. The kit typically includes T7 RNA polymerase, NTPs, buffer, and optional DNase I for template removal. The RNA product can be used directly (after cleanup) for downstream applications (e.g. qRT-PCR, in vitro translation, transfections). In this protocol, we outline the steps using the HiScribe kit: template preparation, transcription reaction setup, DNase treatment (if applicable), RNA purification (if desired), and quality control checks.

1. Prepare all required reagents and consumables:
   • HiScribe T7 High Yield RNA Synthesis Kit (NEB)
   • Linearized DNA template containing a T7 promoter (purified, appropriate concentration)
   • Nuclease-free water
   • DNase I
   • Reaction tubes (RNase-free)
   • Pipettes and RNase-free filter tips
   • RNase inhibitor
   • RNA cleanup kit or reagents (e.g. spin columns, LiCl precipitation, phenol:chloroform, ethanol precipitation)
   • RNAse-free microcentrifuge
   • RNase-free tubes for storage
   • Quantification equipment: NanoDrop
   • Agarose gel electrophoresis
   • Ice bucket, gloves, RNase-free gloves, RNase-away reagent, etc.
2. Template preparation
   • Ensure DNA template is linear (avoid circular plasmid for run-off transcription).
   • Purify the DNA (remove proteins, residual salts). Determine concentration and check purity (A260/A280).
3. Set up transcription reaction
   • Thaw the necessary components at room temperature. Keep the T7 RNA Polymerase Mix on ice.
   • Mix and pulse-spin in a microfuge to collect the solutions to the bottom of the tubes.
   • Set up the reaction at room temperature in the order listed in the table below:

   Components	20 μl reaction	Final amount
   Nuclease-free water	X μL
   10X T7 Reaction Buffer	2 μL	1x
   100 mM ATP	2 μL	10 mM
   100 mM GTP	2 μL	10 mM
   100 mM UTP	2 μL	10 mM
   100 mM CTP	2 μL	10 mM
   Linear Template DNA	X μL	1 μg
   DTT (0.1M)	1 μL	5 mM
   T7 RNA Polymerase Mix	2 μL

4. Incubation / transcription
   • Incubate at 37°C for the duration 2 hours
5. DNase treatment
   • Add DNase I to degrade the DNA template
   • Incubate for the recommended time/temperature to ensure DNA removal.
   • RNA purification (optional but recommended)
6. Quality control and quantification
   • Measure RNA concentration (e.g. NanoDrop).
   • Assess purity (A260/A280, A260/A230).
   • Check integrity and size (agarose gel).
7. Storage
   • Aliquot RNA and store at −80°C (avoid repeated freeze-thaw).
   • Optionally, add RNase inhibitor or buffer stabilizer if needed.

This protocol describes methods for PCR using Q5 High-Fidelity DNA Polymerase, which offers high fidelity (~280X higher than Taq), resulting in ultra-low error rates. Please note that protocols with Q5 High-Fidelity DNA Polymerase may differ from protocols with other polymerases. The conditions recommended below should be used for optimal performance.

1. Prepare all required reagents and consumables:
   • 5X Q5 Reaction Buffer
   • 10mM dNTPs
   • 10μM Forward Primer
   • 10μM Reverse Primer
   • Q5 High-Fidelity DNA Polymerase
   • 5X Q5 High GC Enhancer (optional)
   • Nuclease-Free Water
2. Procedure
   • Assemble all reaction components on ice. Each component should be gently mixed before adding to the reaction in a sterile thin-walled PCR tube. The Q5 High- Fidelity DNA Polymerase may be diluted in 1X Q5 Reaction Buffer just prior to use to reduce pipetting errors. The entire reaction should be mixed again to ensure homogeneous, consistent mixture. Collect all liquid to the bottom of the tube with a quick centrifuge spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid.
   • Quickly transfer the reactions to a thermocycler preheated to the denaturation temperature (98°C) and begin thermocycling. According to the table 1 below.

   Component	25 μl reaction	50 μl reaction
   5X Q5 Reaction Buffer	5 μl	10 μl
   10 mM dNTPs	0.5 μl	1 μl
   10 μM Forward Primer	1.25 μl	2.5 μl
   10 μM Reverse Primer	1.25 μl	2.5 μl
   Template DNA	variable	variable
   Q5 High-Fidelity DNA Polymerase	0.25 μl	0.5 μl
   5X Q5 High GC Enhancer (optional)	(5 μl)	(10 μl)
   Nuclease-Free Water	to 25 μl	to 50 μl

   • Thermocycling Conditions for a Routine PCR:

   Step	Temp
   Initial denaturation	98°C	10 μl
   25–35 Cycles	98°C, 50–72°C, 72°C	5–10 seconds, 10–30 seconds, 20–30 seconds/kb
   Final Extension	72°C	2 minutes
   Hold	4–10°C	∞

Quantitative Reverse Transcription PCR (RT-qPCR) is a sensitive and specific technique used to detect and quantify RNA levels by first converting RNA into complementary DNA (cDNA) using reverse transcriptase, followed by amplification and real-time quantification of specific DNA targets during PCR cycles. RT-qPCR is widely applied in gene expression analysis, pathogen detection, and validation of RNA interference studies. Fluorescent chemistries such as DNA-binding dyes (e.g., SYBR Green) or hydrolysis probes (e.g., TaqMan probes) enable real-time monitoring of amplification, allowing quantification based on fluorescence intensity correlated to the amount of target nucleic acid.

1. Prepare all required reagents and consumables:
   • Luna Universal One-Step Reaction Mix (2X)
   • Luna WarmStart RT Enzyme Mix (20X)
   • Forward primer (10μM) specific to target gene
   • Reverse primer (10μM) specific to target gene
   • RNA samples (purified total RNA, ≤1μg per reaction)
   • Nuclease-free water
   • qPCR tubes or optical 96/384-well plates with sealing film or caps
   • Real-time PCR instrument with SYBR or FAM detection capability
   • Pipettes and sterile filtered tips
2. RNA Preparation
   • Extract and purify RNA from samples using appropriate methods.
   • Quantify RNA concentration by measuring OD260.
   • Prepare RNA dilutions fresh before use, diluting in nuclease-free water or TE buffer if needed.
3. Reaction Setup (20μL total volume)
   • Thaw Luna Universal One-Step Reaction Mix and Luna WarmStart RT Enzyme Mix at room temperature, then place on ice. Mix gently.
   • Prepare a master mix for all reactions (including 10% extra volume) containing:
     • 10μL Luna Universal One-Step Reaction Mix (2X)
     • 1μL Luna WarmStart RT Enzyme Mix (20X)
     • 0.8μL Forward primer (10μM) (final 0.4μM)
     • 0.8μL Reverse primer (10μM) (final 0.4μM)
     • Nuclease-free water to adjust volume before adding RNA template
   • Aliquot 18–19μL of the master mix into each qPCR tube or well.
   • Add 1–2μL of RNA template (≤1μg total RNA) to each tube/well.
   • Seal tubes or plate with optical caps or film.
   • Briefly centrifuge to remove bubbles and collect liquid at the bottom (1 min at 2,500–3,000 rpm)
4. Thermal Cycling Conditions
   • Program the real-time PCR instrument as follows:

   Step	Temperature	Time	Cycles
   Reverse transcription	55°C	10 minutes	1
   Initial denaturation	95°C	1 minute	1
   Denaturation	95°C	10 seconds	40-45
   Annealing/Extenstion	60°C	30 seconds* (+ plate read)	40-45
   Melt Curve (optional)	60-95°C	Instrument-specific	1

5. Data Collection and analysis
   • Use SYBR or SYBR/FAM detection mode on the instrument.
   • Collect fluorescence data at the end of each annealing/extension step.
   • Analyze Cq values and perform relative or absolute quantification as needed.
   • Analyze Cq values and perform relative or absolute quantification as needed.
6. Notes
   • Avoid reverse transcription temperatures below 50°C to ensure full WarmStart RT activation and optimal performance.
   • Primer concentrations and annealing temperatures can be optimized for specific targets.
   • The one-step format reduces contamination risk and simplifies workflow compared to separate RT and qPCR steps.
   • For multiplex assays or probe-based detection, use compatible Luna probe kits and adjust reaction components accordingly.
   • This protocol provides an efficient and reliable method for RNA quantification using the Luna Universal One-Step RT-qPCR Kit, suitable for a wide range of applications including gene expression analysis and pathogen detection

This protocol describes how to properly resuspend lyophilized DNA oligonucleotide primers supplied by Genomed to a final concentration of 100 μM using 10 mM Tris buffer, pH 7.5. The protocol includes preparation of the resuspension buffer and step-by-step instructions for dissolving the primers.

1. Prepare all required reagents and consumables:
   • Lyophilized DNA oligonucleotide primers (ordered from Genomed)
   • Tris base (molecular biology grade)
   • Concentrated HCl (for pH adjustment)
   • Nuclease-free water
   • pH meter or pH indicator strips
   • 0.22 μm sterile filter (for buffer sterilization)
   • Sterile microcentrifuge tubes
   • Pipettes and sterile tips
2. Prepare 1 M Tris-HCl stock solution:
   • Dissolve 12.1 g Tris base in ~80 mL nuclease-free water.
   • Slowly add concentrated HCl while monitoring pH, until pH reaches 7.5.
   • Allow the solution to cool to room temperature, then adjust the pH again if needed.
   • Bring the final volume to 100 mL with nuclease-free water.
   • Sterilize the solution by passing it through a 0.22 μm filter.
3. Prepare 10 mM Tris buffer
   • Dilute the 1 M Tris-HCl stock 1:100 with nuclease-free water (e.g., 1 mL 1 M Tris-HCl + 99 mL water).
4. Resuspension of Lyophilized Primers
   • Determine the required volume. Refer to the information provided by Genomed specifying the volume needed to achieve 100 μM concentration for your primer batch.
   • Add buffer. Add the calculated volume of 10 mM Tris, pH 7.5, directly to the lyophilized primer in its tube.
   • Mix thoroughly. Vortex gently or pipette up and down to ensure complete dissolution. If the primer does not fully dissolve, briefly centrifuge and incubate at room temperature for 10–15 minutes, then mix again.
   • Label and store. Clearly label the tube with primer name, concentration, and date. Store the resuspended primer at –20°C for long-term storage.

The procedure for purifying DNA products from an enzymatic reaction involves adding a DNA-binding buffer to the sample, then applying the mixture to a silica column, where the DNA selectively adsorbs. Subsequent steps include multiple wash cycles to remove residual reagents, followed by elution of the purified DNA using elution buffer or DEPC-treated water. The resulting DNA is inhibitor-free and ready for downstream applications such as ligation, PCR, sequencing, or cloning.

1. Prepare all required reagents and consumables:
   • Clean-Up Concentrator Kit from A&A Biotechnology
   • Microcolumns
   • 1.5 mL elution tubes
   • GI binding solution
   • A1 wash solution
   • Sodium acetate (3 M, ph 5.5)
   • Tris buffer (10 mM, pH 8.5)
   • Additional equipment and reagents
   • 1.5 mL sterile Eppendorf tubes
   • Vortex
   • Microcentrifuge
2. Mix DNA samples (up to 100 μL) with 5 volumes of GI binding solution. Mix the samples by inverting the tubes or vortexing. GI binding solution contains the color pH indicator. Upon mixing the DNA sample with GI binding solution, yellow color of the mixture indicates an optimal pH for DNA binding. If the mixture color turns pink the pH of the solution is too high. In such conditions DNA binds ineffectively to the silica membranes and may be lost. Too high pH can be corrected by adding 1-10 µL of 3 M sodium acetate solution (pH 5.5) (included) and mix. Purification can be continued after reaching a yellow color. optimal condition pH ≤ 7.2 too high pH.
3. Briefly centrifuge the samples to remove the leftovers of solution from the tube walls and caps.
4. Apply samples onto the microcolumns. Close the tubes with the caps.
5. Centrifuge for 30-60 s at 10 000-15 000 RPM.
6. Remove the microcolumns, discard the filtrates. Place the microcolumns to the same tubes.
7. Add 300 µL of A1 wash solution. Close the tubes with the caps.
8. Centrifuge for 30-60 s at 10 000-15 000 RPM.
9. Add 200 µL of A1 wash solution. Close the tubes with the caps.
10. Centrifuge for 2 min at 10 000-15 000 RPM.
11. Transfer the microcolumns to new 1.5 mL tubes (not included).
12. Add 15-30 µL of Tris buffer directly onto the microcolumn resin. Close the tubes with the caps. Applying Tris buffer onto the column be sure that liquid is applied directly onto the resin. If some of the liquid stays on the column wall the elution will be less effective. Elution in a smaller volume is less efficient, but the extracted DNA has a higher concentration. Elution in 30 µL is recommended for fragments over 2000 bp.
13. Incubate for 3 min at room temp.
14. Centrifuge for 2 min at 10 000-15 000 RPM.
15. Remove the microcolumns, close the tubes. Store the tubes with purified DNA at 4-8°C until later use. Elution tube has a long elastic cap connector. It's important to start closing the tube by carefully pressing the cap on the connector side. A "click" - sound confirms proper closure. Different ways of closing may cause opening of the tube during storage.

The NanoDrop is a UV–Vis spectrophotometer that employs surface tension technology to rapidly and accurately assess nucleic acid concentration and purity without requiring cuvettes.

1. Prepare all required reagents and consumables:
   • Autoclaved dH2O
   • Fiber-free tissues
   • NanoDrop spectrophotometer
   • Pipettes and tips
   • Samples
2. Start the NanoDrop spectrophotometer.
3. Select the DNA measurement ‘Nucleic Acid’ in the NanoDrop menu for examining DNA sample.
4. Clean the surface of the NanoDrop with dH2O and a fiber free-tissue.
5. Perform a calibration and blank measurement by entering one drop of 1 μL autoclaved dH2O.
6. Clean the surface again and place 1 μL per sample on the NanoDrop and measure the concentration.

DNA electrophoresis is a technique used to separate fragments of deoxyribonucleic acid (DNA) in an electric field based on their size. Due to the negative charge of phosphate groups in the DNA backbone, the molecules migrate towards the anode. Smaller fragments move more rapidly through the polymer network of the gel than larger ones, which allows for the assessment of fragment length, verification of PCR or cloning quality, and the isolation of specific fragments for further analyses.

1. Prepare all required reagents and consumables:
2. Reagents
   • Agarose
   • 1X TBE Buffer
   • GelRed nucleic acid stain
   • 6X Loading dye
   • DNA Ladder (e.g. 1 kb Plus Ladder)
   • Samples (PCR product or purified DNA)
3. Equipment
   • Electrophoresis chamber with power supply
   • Gel casting trays with combs
   • Casting stands
   • Microwave oven
   • UV lamp or blue-light transilluminator for gel visualization
4. Gel preparation
   • Calculate the gel volume and percentage based on the number and characteristics of the samples.
   • Use an analytical balance to weigh the appropriate amount of agarose (e.g., 1 g of agarose per 100 mL of 1X TBE for a 1 % gel).
   • Pour 1X TBE buffer into a flask in a volume equal to the final gel volume, then add the agarose.
   • Heat the mixture in a microwave until the agarose is completely dissolved. Remove every 30 seconds to swirl gently and verify that a clear solution has formed.
   • Cool the gel solution under running tap water until it reaches approximately 55°C.
   • Add the recommended amount of GelRed according to the manufacturer’s instructions, and mix gently.
   • Attach casting stands to the gel tray, insert the comb, pour in the gel solution, and allow it to solidify at room temperature for 20–30 minutes.
   • Once polymerized, remove the casting stands and carefully extract the comb to create the sample wells.
5. Sample Treatment
   • Prepare DNA samples and ladders in Eppendorf tubes.
   • Add loading dye to each sample at a ratio of 1 part loading dye to 5 parts sample (e.g., 5 μL sample + 1 μL loading dye).
   • Gently mix by pipetting, avoiding the introduction of air bubbles.
   • Briefly centrifuge the samples (1–2 s) to collect all contents at the bottom of the tube.
6. Electrophoresis & Analysis
   • Place the concentrated gel in the electrophoresis chamber and add 1X TBE buffer until it covers the gel (approximately 3–5 mm above the surface).
   • Carefully load the ladder and samples into the wells, loading the ladder first, followed by the samples.
   • Connect the electrodes and set the voltage to 5–10 V/cm (for example, for a 10 cm gel, set to 100 V).
   • Start the power supply and run the electrophoresis for the appropriate duration (usually 30–60 minutes) until the tracking dyes have migrated to the desired distance from the origin.
   • After completion, turn off the power supply, remove the gel, and place it on a transilluminator to visualize the DNA fragments.

The DNA extraction process from agarose gel involves excising the target band from the gel, dissolving the matrix in a dissolving buffer, and then selectively binding the DNA to a silica column. Subsequent steps include the removal of contaminants through multiple wash steps, followed by elution of the DNA in a small volume of buffer or water. The purified DNA obtained is ready for downstream applications such as ligation, sequencing, cloning, or PCR.

1. Prepare all required reagents and consumables:
2. Gel-Out kit A&A Biotechnology
   • Minicolumns
   • R7SI agarose solution
   • A1 wash solution
   • Sodium acetate (3 M, pH 5.5)
   • Isopropanol
   • TE buffer
3. Additional Required Reagents/Equipment
   • 1.5 mL sterile Eppendorf tubes
   • Incubator or thermoblock set to 50°C
   • Vortex
   • Microcentrifuge
4. Cut out the agarose slices (up to 200 mg) containing DNA. Transfer agarose slices to Eppendorf tubes (not included). Agarose gel electrophoresis can be performed in the presence of either TAE or TBE buffer.
5. Add an appropriate volume of R7SI agarose melting solution:
   • < 2% agarose gel – 400 μL
   • ≥ 2% agarose gel – 500 μL
   Incubate samples at 50°C until complete dissolution of agarose slices. Mix the samples by inverting the tubes or vortexing a few times. Agarose melting solution R7SI contains the color pH indicator. Upon mixing the DNA sample with R7SI agarose melting solution, yellow color of the mixture indicates an optimal pH for DNA binding. If the mixture color turns pink the pH of the solution is too high. In such conditions DNA binds ineffectively to the silica membranes and may be lost. Too high pH can be corrected by adding 1–10 μL of 3 M sodium acetate solution (pH 5.5) (included) and mix. Purification can be continued after reaching a yellow color. optimal condition pH ≤ 7.2 too high pH
6. Add an appropriate volume of isopropanol:
   • < 2% agarose gel – 200 μL
   • ≥ 2% agarose gel – 250 μL
   Mix by inverting the tubes.
7. Briefly centrifuge the samples to remove the leftovers of solution from the tube walls and caps.
8. Apply samples onto the minicolumns.
9. Centrifuge for 30 s at 10 000–15 000 RPM.
10. Remove the minicolumns, discard the filtrate. Place the minicolumns to the same tubes.
11. Add 600 μL of A1 wash solution.
12. Centrifuge for 30 s at 10 000–15 000 RPM.
13. Remove the minicolumns, discard the filtrate. Place the minicolumns to the same tubes.
14. Add 300 μL of A1 wash solution.
15. Centrifuge for 1 min at 10 000–15 000 RPM.
16. Remove the minicolumns, discard the filtrate. Place the minicolumns to the same tubes.
17. Centrifuge for 1 min at 10 000–15 000 RPM.
18. Transfer the minicolumns to new 1.5 mL tubes.
19. Add 50 μL of TE buffer or sterile water (not included) directly onto the minicolumn resin. Applying elution liquid (TE buffer or sterile water) onto the minicolumn be sure that liquid is applied directly onto the resin. If some of the liquid stays on the column wall the elution will be less effective. Elution in a smaller volume is less efficient, but the extracted DNA has a higher concentration. Elution in 50 μL volume is more efficient, but DNA has a lower concentration.
20. Incubate for 3 min at room temp.
21. Centrifuge for 1 min at 10 000–15 000 RPM.
22. Remove the minicolumns and store the tubes with purified DNA at 4–8°C until later use.

Isolation of plasmids using the Midi‐prep method is a process for purifying plasmid DNA from a large volume of bacterial culture via kits equipped with ion‐exchange columns. This technique relies on alkaline lysis of the cells, followed by selective binding of plasmid DNA to the column matrix, washing away contaminants, and eluting the pure DNA, which is then suitable for downstream applications such as transfection, PCR, or sequencing.

1. Prepare all required reagents and consumables:
   • Kit Components (per 1 isolation)
   • Plasmid 200 column (1x)
   • Filtration column (1x)
   • 50 mL tubes (2x)
   • L1 cell suspension solution (5 mL)
   • L2 lysis solution (5 mL)
   • L3 neutralizing solution (5 mL)
   • LPS-out endotoxin removal solution (15mL)
   • K2P wash solution (20 mL)
   • K3 elution solution (6 mL)
   • Precipitation enhancer (25 μL)
   • Isopropanol (5 mL)
   • TE buffer (0.2–1 mL for resuspension)
   • Additional Required Reagents/Equipment
   • 70% Ethanol (2 mL)
   • Sterile 15 mL and 50 mL Falcon tubes
   • Microcentrifuge capable of 11,000 × g
   • Sterile water (optional)
   • Pipettes and tips
   • Sterile cotton buds (for drying)
   • Personal protective equipment (PPE)
   • Safety notes
   • L2 lysis solution and K3 elution solution contain hazardous chemicals. Refer to the kit’s safety data for handling instructions.
   • Isopropanol is flammable and an irritant-work in a ventilated area and away from ignition sources.
   • Wear gloves, lab coat, and eye protection throughout the procedure.
2. Cell harvesting
   • Centrifuge up to 100 mL of overnight E. coli culture at 4,000–6,000 × g for 10 min.
   • Discard the supernatant.
   • Resuspend the bacterial pellet in 5 mL of L1 cell suspension solution. Note: Solution should change from deep pink to opaque light pink; ensure no pellet remains at the bottom.
3. Cell lysis
   • Add 5 mL of L2 lysis solution.
   • Gently invert the tube several times (do not vortex).
   • Incubate at room temperature for 5 min. Note: Lysate should become clear and raspberry-colored. If not, mix gently and incubate for an additional 3 min.
4. Neutralization
   • Add 5 mL of L3 neutralizing solution. Note: Solution should turn yellowish, indicating complete neutralization.
5. Filtration
   • Transfer the lysate to the filtration column.
   • Centrifuge at 1,500 × g for 5 min.
6. Endotoxin removal
   • Add 1.2 mL LPS-out solution to the filtered lysate. Mix and keep on ice for 30 min.
7. DNA binding
   • Place a Plasmid 200 column into a clean 50 mL tube.
   • Apply the clear filtrate onto the column and allow it to flow through completely.
8. Column Wash
   • Add 20 mL of K2P wash solution to the column.
   • Allow the solution to flow through.
9. Elution
   • Transfer the Plasmid 200 column to a new 50 mL tube.
   • Add 6 mL of K3 elution solution.
   • Collect the eluate.
10. DNA Precipitation
    • Transfer the eluate to a new 15 mL tube.
    • Add 25 μL precipitation enhancer and 5 mL isopropanol. Note: For sensitive applications, omit the enhancer if desired.
    • Mix by gentle inversion.
    • Centrifuge at 11,000 × g for 10 min. Note: A light-blue DNA pellet should be visible.
11. Wash DNA Pellet
    • Carefully discard the supernatant (do not lose the pellet).
    • Add 2 mL of 70% ethanol.
    • Mix and centrifuge at 11,000 × g for 3 min.
12. Dry DNA Pellet
    • Carefully discard the supernatant.
    • Air dry the pellet at room temperature (10 min, tube inverted).
    • Remove any residual ethanol with a sterile cotton bud if necessary.
13. DNA Resuspension
    • Dissolve the DNA pellet in 0.2–1 mL of TE buffer or sterile nuclease-free water. Note: The blue color aids in visual confirmation of dissolution.
14. Storage
    • Store the purified plasmid DNA at 4–8°C.
15. Color Indicators (LySee System)
    • L1 addition: Deep pink → opaque light pink (complete resuspension)
    • L2 addition: Opaque light pink → clear raspberry (complete lysis)
    • L3 addition: Raspberry → yellowish (complete neutralization)
16. Troubleshooting
    • L2 solution cloudy: Warm at 40°C until clear before use.
    • Pellet loss: Always pour supernatant into a separate tube to recover pellet if accidentally lost.
    • Incomplete resuspension/lysis/neutralization: Confirm color changes at each step.
17. Waste Disposal
    • Dispose of all hazardous liquids and biological waste according to institutional guidelines.

Isolation of plasmids using the Mini-prep method is a process for purifying plasmid DNA from a small volume of bacterial culture. This technique relies on alkaline lysis of bacterial cells, which causes denaturation of genomic DNA and cellular proteins, while small, circular plasmid DNA retains its structure. Following lysis, contaminant precipitation occurs and selective recovery of plasmid DNA is achieved through binding to the column matrix, washing, and elution of pure DNA in small volumes. Plasmids obtained through such isolation can be applied in processes such as bacterial transformations, PCR reactions, or sequencing.

1. Prepare all required reagents and consumables:
   • Centrifuge
   • Sterile 1.5 mL Eppendorf-type tubes
   • Sterile nuclease-free water
   • Plasmid Mini Kit A&A Biotechnology
   • Microcolumns
   • L1 Cell Resuspension Solution
   • L2 Lysis Solution
   • GL3 Neutralization Solution
   • W First Wash Solution
   • A1 Second Wash Solution
   • TE Buffer
2. Centrifuge 1.5–3 mL of overnight bacterial culture in 12 500 RCF for 10 minutes.
3. Remove the supernatant and thoroughly resuspend the pellet in 200 μL of L1 Cell Resuspension Solution. Note: During resuspension of the bacterial pellet, the solution will change in appearance from completely transparent with a dark pink tint to opaque with a light pink tint. Resuspension can be considered complete once the pellet at the bottom of the tube has completely disappeared.
4. Add 200 μL of L2 Lysis Solution and gently mix until complete lysis occurs. Note: After adding L2 Lysis Solution, gently mix the tube contents to avoid chromosomal DNA fragmentation. Several gentle tube inversions are typically sufficient. The mixture should change in appearance and coloration.
5. Incubate for 3 minutes at room temperature. Note: After 3 minutes of incubation, the lysate should be completely clear and uniformly pink. If not, mix the lysate several times and extend incubation for an additional 3 minutes.
6. Add 400 μL of GL3 Neutralization Solution and gently mix until the pink coloration of the lysate disappears. Note: Upon addition of GL3 Neutralization Solution, rapid precipitation of potassium SDS salts, chromosomal DNA, and some proteins occurs. After mixing, the tube contents should change to a light yellow color. The absence of pink coloration indicates complete neutralization and successful completion of alkaline lysis.
7. Centrifuge the lysate for 10 minutes at 12,500 RCF.
8. Carefully apply the lysate (supernatant) to the minicolumn.
9. Centrifuge for 1 minute at 12,500 RCF.
10. Remove minicolumns from tubes, discard the flow-through, and replace minicolumns in the same tubes.
11. Add 500 μL of first wash solution W.
12. Centrifuge for 1 minute at 12,500 RCF.
13. Remove minicolumns from tubes, discard the flow-through, and replace minicolumns in the same tubes.
14. Add 600 μL of second wash solution A1.
15. Centrifuge for 2 minutes at 12,500 RCF.
16. Transfer minicolumns to new 1.5 mL tubes.
17. Apply 60 μL of TE buffer or sterile water to the column matrix.
18. Incubate samples for 3 minutes at room temperature.
19. Centrifuge for 1 minute at 12,500 RCF.
20. Remove minicolumns and store plasmid DNA at 4-8°C until further analysis.

The NanoDrop is a UV–Vis spectrophotometer that employs surface tension technology to rapidly and accurately assess nucleic acid concentration and purity without requiring cuvettes.

1. Prepare all required reagents and consumables:
   • Autoclaved dH2O
   • Fiber-free tissues
   • NanoDrop spectrophotometer
   • Pipettes and tips
   • Samples
2. Start the NanoDrop spectrophotometer.
3. Select the RNA measurement ‘Nucleic Acid’ in the NanoDrop menu for examining RNA sample.
4. Clean the surface of the NanoDrop with dH2O and a fiber free-tissue.
5. Perform a calibration and blank measurement by entering one drop of 1 μL autoclaved dH2O.
6. Clean the surface again and place 1 μL per sample on the NanoDrop and measure the concentration.

The total RNA was isolated using a protocol involving cell lysis in a phenol–guanidine buffer, followed by specific binding of RNA to a silica column. The purified RNA can be employed, among other applications, for cDNA synthesis and RT-qPCR analysis.

1. Prepare all required reagents and consumables:
   • Total RNA Mini Kit
   • Minicolumns – 100 pcs
   • 2 mL Tubes – 100 pcs
   • A1 Washing Solution – 200 mL
   • Phenol – 100 mL
   • Isopropanol – 30 mL
   • Ultrapure Water – 15 mL
   • Sterile 1.5 mL Eppendorf-type tubes
   • Chloroform
   • Microcentrifuge
   • Incubator at 50°C
2. Preparation of Cell Culture Material
   • Centrifuge cells (1×10⁶) and remove the supernatant.
3. RNA Isolation
   • Add 800 μL of Fenozol (phenol-based solution) to the prepared sample. Mix by pipetting until complete cell lysis. Notes: Fenozol inactivates endogenous RNases. Samples in Fenozol can be stored:
     • Up to 1 year at -80°C
     • Up to 1 week at 2–8°C
     • Up to 24 hours at room temperature
     • ⚠️ Fenozol contains phenol. Avoid skin contact; wear protective gloves.
   • Incubate the sample for 5 minutes at 50°C.
   • Add 200 μL chloroform (not included in the kit) to the lysate. Gently mix by inverting the tube several times.
   • Let the sample sit for 3 minutes at room temperature (20–25°C).
   • Centrifuge for 10 minutes at 10,000–12,000 RPM.
   • Transfer the upper aqueous phase (~450 μL supernatant) to a new 1.5 mL tube.
   • Add 250 μL isopropanol and mix thoroughly.
   • Apply the mixture to a minicolumn.
   • Centrifuge for 1 minute at 10,000–12,000 RPM.
   • Transfer the minicolumn to a new 2 mL tube.
   • Add 700 μL Wash Solution A1.
   • Centrifuge for 1 minute at 10,000–12,000 RPM.
   • Discard the flow-through and reinsert the minicolumn into the same tube.
   • Add 700 μL Wash Solution A1 and centrifuge again for 1 minute at 10,000–12,000 RPM.
   • Discard the flow-through and reinsert the minicolumn.
   • Add 200 μL Wash Solution A1 and centrifuge for 2 minutes at 10,000–12,000 RPM.
   • Transfer the minicolumn to a new 1.5 mL tube.
   • Add 100 μL ultrapure water directly to the center of the minicolumn membrane.
   • Let the column sit for 3 minutes at room temperature.
   • Centrifuge for 1 minute at 10,000–12,000 RPM.
   • Discard the minicolumn. Store the purified RNA in the tube at -20°C or -80°C until further analysis.