NOTEBOOK

（Wet Lab Notebook）

Main Timeline:

（Each month is depicted with a grid system where each grid corresponds to 3 days. The shaded regions indicate the days designated for work.）

January:

Establishment of the project team Starting from 11th				Starting from 11th
Development of Project Plan				Starting from 11th
Recruiting of new members				Starting from 11th

February:

Establishment of the project team
Development of Project Plan
Recruiting of new members

March:

Establishment of the project team
Development of Project Plan

April:

Establishment of the project team
Development of Project Plan

May:

Subject determination
Theoretical study

June:

Theoretical study
Subject determination
Experimental design
Safety training
Experimental design

July:

Experimental design
Problems discussion
Record experiment report
Building the Team Wiki

August:

Implement experimental plan
Problems discussion
Analysis of experimental results
Building the Team Wiki

September:

Problems discussion
Building the Team Wiki
Education
Preparation of defense materials
Analysis of experimental results

October:

Preparation of defense materials
Education
Project review
Building the Team Wiki

Record of Main Experimental Content

Pre-experiment safety training:

The main content is as follows:

I. Always ensure to wear the necessary personal protective equipment, including lab coats, gloves, and safety goggles, when working in the laboratory.

II. Be familiar with the location and proper usage of emergency equipment, such as eyewash stations, fire extinguishers, and emergency showers.

III. Refrain from consuming food or beverages or applying cosmetics in the laboratory to avoid accidental exposure to hazardous substances.

IV. Adhere to correct procedures for handling and disposing of chemicals, biohazardous materials, and sharps to reduce potential risks.

V. Maintain cleanliness and organization in work areas to prevent accidents and the spread of contaminants.

VI. Promptly report any spills, accidents, or injuries to the laboratory supervisor.

VII. Refrain from conducting any unauthorized activities or experiments without appropriate training and supervision.

VIII. Utilize equipment and instruments solely for their designated purposes and operate them following safety protocols.

IX. Acquaint yourself with emergency protocols, including evacuation routes and assembly points, in the event of emergencies.

X. Attend regular laboratory safety training sessions and strictly adhere to safety protocols and guidelines to ensure a secure working environment for both yourself and others.

Part 1:

I: Amplify the target gene by PCR, verify band size through agarose gel electrophoresis, recover the target gene from the gel, and store it at -20°C for future use.

II: Heat shock the recombinant product into Escherichia coli DH5a, plate on antibiotic-resistant agar plates, and incubate at 37°C.

III: Inoculate the shaken culture with X-3 and XI-2 plasmid backbones and culture overnight at 37°C.

Detailed records of the main experiments:

I: The basic steps of PCR technology for amplifying the target gene include denaturation, annealing, and extension. Denaturation: Initially, DNA is heated to 90-95°C, breaking the hydrogen bonds of the double-stranded DNA template to form single-stranded DNA. This step prepares for the subsequent annealing and extension processes. Annealing: Subsequently, the temperature is lowered to 55-60°C, allowing the primers to bind to the single-stranded DNA template, forming localized double strands. This step utilizes the principle of base pairing to facilitate the specific binding of primers to the target region of the DNA template. Extension: Finally, the temperature is raised again to 70-75°C, where Taq polymerase (optimal activity around 72°C) utilizes dNTPs to extend from the 5′ to 3′ end of the primer, synthesizing a complementary DNA strand to the template. This step completes the synthesis of a new DNA strand, thereby achieving DNA replication. Each cycle involves denaturation, annealing, and extension, doubling the DNA content. By repeating this process multiple times (usually 25-30 cycles), specific amplification of a particular DNA fragment can be achieved. This PCR process is based on the fundamental principle of DNA replication, allowing for the extensive replication of the target DNA sequence through repetitive cycling to meet subsequent experimental or application needs.

II: Preparation of medium Yeast extract peptone dextrose YPD/X medium for yeast cultivation. The concentrations of yeast extract and peptone are 10 g/L and 20 g/L, respectively, and they are separately sterilized with 400 g/L glucose or xylose solutions. After sterilization, the calculated glucose or xylose solution is added to YP to achieve a final concentration of 20 g/L. The preparation of YPD/X solid medium only requires the addition of 2% (w/v) agar during YP sterilization.

Part 2:

I: Verify pscm-ISU1 and pscm-nfsI colonies by colony PCR and confirm target bands through agarose gel electrophoresis.

II: Perform overlap PCR amplification of TEF1-XI-ADH1 and GAP-XI-CYC1, verify band sizes through agarose gel electrophoresis, and proceed with DNA gel extraction.

III: Extract X-3 and XI-2 backbone plasmids, perform double enzyme digestion with XhoI and MssI, and proceed with DNA gel extraction.

IV: T4 ligation of TEF1-XI-ADH1 with X-3 and TEF1-XI-ADH1 with XI-2, incubate at 37°C for 2-3 hours.

V: Heat shock the recombinant products X-3-XI and XI-2-XI into Escherichia coli DH5a, plate on Amp plates, and culture overnight at 37°C.

VI: Shake-culture X-3-XI and XI-2-XI strains.

Detailed records of the main experiments:

Colony PCR involves several key steps, including primer design, colony picking, PCR reaction, and result analysis.

Primer Design: The first step in colony PCR is primer design, which includes three strategies: insert-specific primers, vector-specific primers, and directional-specific primers. Primer design is crucial in colony PCR as it directly impacts the specificity and efficiency of the PCR reaction. Insert-specific primers are annealed to the insert-specific sequence, vector-specific primers are annealed to the vector sites flanking the insert, and directional-specific primers determine the orientation of the insert. Before screening colonies with colony PCR primers, it is essential to test them. One recommended method is to use the target fragment primers on one end and validation primers on the vector backbone on the other end to confirm the successful construction of the fusion vector.

Colony Picking: Use a sterile toothpick or pipette tip to pick a single colony and spot it on a selective plate for clonal preservation (labelling). Then, place the toothpick or pipette tip with the colony into a PCR tube containing the PCR mix to release the cells into the PCR mix as a template.

PCR Reaction: Put the PCR mix containing the bacterial cells into a PCR machine and amplify following standard conditions. The PCR reaction typically includes Taq buffer, dNTPs, forward and reverse primers, polymerase, etc., with specific components and concentrations adjusted based on experimental requirements and optimization conditions.

Result Analysis: After PCR amplification, analyze the PCR products using agarose gel electrophoresis to visualize the target bands. If the band size matches the expected size, it indicates the presence of the target gene or plasmid in the colony, confirming a positive clone.

Additionally, sterile techniques, contamination prevention, primer specificity, and efficiency considerations in primer design are crucial in colony PCR to ensure the accuracy and reliability of experimental results.

Part 3:

I: Verify X-3-XI and XI-2-XI colonies through colony PCR and confirm band sizes by agarose gel electrophoresis.

II: Extract X-3-XI and XI-2-XI plasmids, perform double enzyme digestion on the plasmids, and recover the target bands from the gel.

III: Perform T4 ligation of the vector and target gene fragments, followed by heat shock transformation into Escherichia coli.

IV: Shake-culture the phcas9 and gRNA strains.

V: Prepare YPD medium for bacterial culture, sterilize, and set aside.

Detailed records of the main experiments:

The key steps in microbial inoculation and cultivation typically include the following: Sterilization: First, all tools and containers used need to undergo sterilization to prevent contamination by external microorganisms. This includes inoculation loops, needles, culture media, and petri dishes.

Inoculation: After sterilization, a small amount of the original microbial culture is transferred using an inoculation loop or needle and then inoculated onto the culture medium. There are various inoculation methods, such as streaking and spread plating, with the specific method depending on the experimental purpose and microbial characteristics.

Cultivation: After inoculation, the petri dishes are placed at the appropriate temperature for cultivation. Most microorganisms are typically cultivated at 37°C for 18-24 hours, but the exact temperature and time may vary depending on the type of microorganism.

Observation and Recording: During cultivation, it is important to regularly observe the growth of microorganisms, including the morphology, color, and size of colonies, and record these observations. This helps in subsequent identification and applications.

Throughout the entire process, the aseptic technique is crucial, as even minor contamination can lead to experimental failure. Therefore, strict adherence to aseptic techniques from sterilization to vaccination is essential.

Part 4:

I: Verify X-3-2XI and XI-2-2XI colonies through colony PCR and confirm band sizes by agarose gel electrophoresis.

II: Extract plasmids: X-3-2XI and XI-2-2XI, pHcsa9 and gRNA, stored at -20°C for future use.

III: Perform single enzyme digestion with NotI on X-3-2XI and XI-2-2XI plasmids, confirm band sizes by agarose gel electrophoresis, and recover bands after enzyme digestion.

IV: Shake-culture the following strains: xyl-cas9, Xyl, xyl-8XI, xyl-8XI-nfsI, xyl-8XI-ISU1, xyl-8XI-nfsI-ISU1.

Detailed records of the main experiments:

The basic principle of enzyme digestion and ligation:

Enzyme digestion and ligation technology relies on the action of DNA restriction endonucleases. These enzymes can specifically recognise particular sites within DNA sequences and cleave the DNA at these sites, generating either sticky ends or blunt ends. By using restriction endonucleases, the target gene and vector DNA can be cut into linear molecules, which are then ligated together using DNA ligase to form a recombinant plasmid.

Specific steps of enzyme digestion and ligation:

Vector preparation: Select an appropriate vector, such as a plasmid, that contains an origin of replication, selection markers, and multiple cloning sites. Use restriction endonucleases to cut the vector, generating linearised vector DNA.

Insert preparation: Similarly, restriction endonucleases are used to cut the target gene, generating a linearised fragment of the target gene DNA.

Ligation reaction: Purify the digested vector and target gene fragment and use enzymes like T4 DNA ligase to ligate them together. The ligation reaction requires the appropriate reaction conditions, such as ATP, DTT, and Mg2+.

Transformation and selection: Transform the ligation product into competent cells and select for successfully transformed clones using antibiotic resistance or other selection markers.

Through these steps, the target gene can be stably inserted into the vector, creating a recombinant plasmid, which serves as the basis for subsequent gene expression, functional studies, and other applications.

Part 5:

I: After transferring and culturing the yeast, culture at 30 degrees Celsius until reaching an OD of 0.8-1.0 to prepare yeast-competent cells.

II: Transform the pHcsa9 plasmid into xyl yeast using the LiAc method, and transform both the gRNA and the recovered two-copy fragments into xyl-cas9 yeast.

III: Following heat shock transformation, recover by incubating for 1-2 hours, plate on YPD-Amp plates, and culture at 30 degrees Celsius overnight.

Detailed records of the main experiments:

The preparation steps for the yeast competent cells mainly include the following stages:

Dip a sterile toothpick into an appropriate amount of yeast glycerol stock and streak a line on a YPD solid agar plate to isolate single colonies. Pick a single colony and inoculate it into a 3 mL YPD liquid culture tube, then incubate overnight at 30°C and 240 rpm on a shaker.

Dilute the overnight culture 10-fold and measure the OD600, ensuring that the OD is more significant than 0.2. Inoculate a 250 mL conical flask containing 25 mL of 2×YPD medium with an initial OD600 of 0.2, and culture at 30°C and 240 rpm until the OD600 reaches 0.8 to 1.0. Centrifuge the culture at 20°C and 3000 rpm for 5 minutes to collect the cells, resuspend the cells in 25 mL of sterile water, and repeat the centrifugation step twice.

Prepare a 42°C water bath and a 100°C metal bath. Add 1 mL of sterile water to resuspend the cells, transfer the cell suspension to a 1.5 mL centrifuge tube, centrifuge at 12000 rpm for 30 seconds, discard the supernatant, collect the cells, and obtain the yeast-competent cells.

The prepared, competent cells should be aliquoted and stored at -80°C for long-term storage and use.

These steps and considerations ensure the successful preparation of brewing yeast competent cells, providing a foundation for subsequent genetic operations and experiments.

Part 6:

I: Verify the xyl-8XI single clone yeast colony by colony PCR and confirm the target bands by agarose gel electrophoresis.

II: Amplify the upstream and downstream homologous arms of the nfs1 gene and recover the fragments.

III: Transform the recovered fragments and the pscm-nfs1 plasmid into the xyl-8XI strain and culture overnight at 30 degrees Celsius.

Detailed records of the main experiments:

S.cerevisiae competent cell transformation method using LiAc.

Table 1 Reagents for LiAc conversion

Compositions	Volume (μL)
PEG 3350（500 g/L）	240
LiAc（1 M）	36
Salmon sperm ssDNA（2 mg/L）	50
DNA + H2O	34

First, place 2 mg/L salmon sperm DNA in a 100°C metal bath for 5 minutes. After 5 minutes, quickly remove it and put it on ice for later use.

Add the following components sequentially to the centrifuge tube containing the collected competent cells, following the order and quantities as shown in Table 1. Perform a 42°C heat shock for 15 minutes, centrifuge at 12000 rpm for 30 seconds, discard the supernatant, add 1 mL YPD liquid medium, resuspend the cells, and seal with parafilm. Place it on a shaker, recover at 30°C and 240 rpm for 2 hours, then centrifuge at 3000 rpm for 4 minutes, discard 600 μL of the supernatant, resuspend, and spread 200 μL of the cell suspension evenly on a solid selection plate. Invert the plate and culture at 30°C for two days.

Part 7:

I: Verify the xyl-8XI-nfs1 single clone yeast colony by colony PCR and confirm the target bands by agarose gel electrophoresis.

II: Amplify the upstream and downstream homologous arms of the ISU1 gene and recover the fragments.

III: Transform the recovered fragments and the pscm-ISU1 plasmid into the xyl-8XI strain and culture overnight at 30 degrees Celsius.

Part 8:

I: Verify the xyl-8XI-ISU1 single clone yeast colony by colony PCR and confirm the target bands by agarose gel electrophoresis.

II: Transform the recovered fragments and the pscm-ISU1 plasmid into the xyl-8XI-Nfs1 strain and culture overnight at 30 degrees Celsius.

III: Inoculate and shake culture the 5 constructed strains overnight after the completion of the transformation.

Part 9:

I: Verify the xyl-8XI-ISU1 single clone yeast colony by colony PCR and confirm the target bands by agarose gel electrophoresis.

II: Transform the recovered fragments and the pscm-ISU1 plasmid into the xyl-8XI-Nfs1 strain and culture overnight at 30 degrees Celsius.

III: Shake the culture of the 5 constructed strains overnight after the completion of the transformation.

Part 10:

Ferment the constructed 4 strains along with the control strain in YPDX medium to assess the xylose metabolism of the strains.