Week 1 Lab: Pipetting

Performing Serial Dilution

Dilution Practice 1

Dilution Practice 2

1. The stock concentration of a mystery substance (MS) is 5 M. If the molar mass of MS is 532 g/mol, what’s the concentration of the stock concentration in g/mL? To make your life easier, you can use one of many online calculators.

The stock concentration of MS is 2.66 g/mL.
Steps:
First, Convert moles to grams. A 5 M solution contains 5 moles per liter: 5 mol/L * 532 g/mol = 2660 g/L
Second, Convert grams per liter to grams per milliliter: 2660 g / 1000 mL = 2.66 g/mL

2. You will perform a serial dilution to get 100 uM of MS. Devise a plan to dilute a 5 M MS solution to 100 uM. How many dilution steps will we need? Which tubes should we use? Which pipettes?

Three dilution steps are needed using P10 & P1000 pipettes with three 1.5 mL microcentrifuge tubes.
Steps:
First, Calculate the total dilution factor.
5 M = 5 mol/L. The target is 100 µM = 1 × 10^-4 mol/L. Total dilution factor = (5 mol/L) / (1 × 10^-4 mol/L) = 50,000.
Second, Break this into three steps.
50,000 = 100 × 100 × 5, so we will do a 1:100 dilution, then another 1:100 dilution, then a 1:5 dilution.
Third, Use three 1.5 mL microcentrifuge tubes (Tube 1, Tube 2, Tube 3).
• Tube 1: 1:100 dilution from the 5 M stock. Add 990 µL diluent + 10 µL of 5 M MS (P1000 + P10).
   Final concentration in Tube 1 = (5 M) / 100 = 0.05 M = 50 mM.
• Tube 2: 1:100 dilution from Tube 1. Add 990 µL diluent + 10 µL from Tube 1 (P1000 + P10).
   Final concentration in Tube 2 = (50 mM) / 100 = 0.5 mM = 500 µM.
• Tube 3: 1:5 dilution from Tube 2. Add 800 µL diluent + 200 µL from Tube 2 (both with P1000).
   Final concentration in Tube 3 = (500 µM) / 5 = 100 µM.<br

3. Fill out the following chart to prepare a final reaction with 60 uL reaction volume. Why did we make 100 uM MS if we actually need 40 uM MS? Why not prepare 40 uM in serial dilutions?

Check with C₁V₁ = C₂V₂:

• Loading dye: (6X) · V₁ = (1X) · (60 µL) → V₁ = 10 µL
• MS: (100 µM) · V₁ = (40 µM) · (60 µL) → V₁ = (40/100) · 60 µL = 24 µL
• dH₂O: 60 µL total − 10 µL dye − 24 µL MS = 26 µL

We made 100 µM MS as a working stock because:

• Pipetting accuracy:
  It’s much easier and more accurate to pipette volumes like 10–25 µL from a 100 µM stock than to pipette tiny volumes from a much more dilute 40 µM stock.
  
• Flexibility:
  A 100 µM stock can be used to make many different final concentrations (e.g., 10, 20, 40, 50 µM) simply by adjusting the volume added. A 40 µM stock would only be useful for reactions requiring exactly 40 µM.
  
• Separation of tasks:
  Preparing a 100 µM intermediate stock allows you to complete the serial dilution step once. After that, setting up reactions becomes a simple mixing step rather than repeating dilution calculations every time.
  
• Cleaner math for the final mix:
  Using a 100 µM stock makes the final reaction calculation simple. Going from 100 µM to 40 µM in a 60 µL reaction is a single C₁V₁ = C₂V₂ step, and it uses pipetting volumes that are accurate and easy to measure.
  

Week 10 Lab: Mass Spectrometry

Figure 1. Chromatography using various solvent.

Instead of Mass Spectrometry, we did a Chromatography instead. Chromatography relies on a substance’s chemical affinity to separate components out of a physical mixture. Mass spectrometry, on the other hand, relies on the mass-to-charge ratio (m/z) of fragmenmted molecules moving through electromagnetic field to identify exact chemical structure.

Figure 2. Diagram Explainations & fellas.

Liquid Chromatography-Mass Spectrometry (LC-MS) While Chromatography and Mass Spectrometry are fundamentally different, laboratories processes might combine them into a single workflow, such as hyphenated techniques. LC‑MS is a hyphenated analytical technique that integrates the separation power of liquid chromatography with the molecular detection capabilities of mass spectrometry. The LC system resolves complex mixtures into individual components, while the MS system ionizes each eluting compound and records its mass‑to‑charge profile. Together, they provide highly sensitive, selective, and structurally informative analysis suitable for applications ranging from small‑molecule quantification to large‑scale proteomics.

Week 12 Lab: Bioproduction

Week 2 Lab: DNA Gel Art

Gel Art - Restriction Digests and Gel Electrophoresis

Protocol | Part 0: Designing your Gel Art

Protocol | Part 1a: Preparing a 1% agarose electrophoresis gel

gel protocals

Protocol | Part 1b: Restriction Digest

Protocol | Part 2: Gel Run

Protocol | Part 3: Imaging Your Results with a Transilluminator

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Week 3 Lab: Opentrons Art

Lab 3 - Opentron Art

Design

Opentrons

Week 6 Lab: Restriction Enzyme Digest I - MiniPrep

Figure 1. Diagrams explaining Gibson Assembly & Golden Gate Assembly by Dr. Andrew Scarpelli at community biolab, ChiTownBio.

For this week, we performed a restriction digest and ligation in lieu of using Gibson Assembly or Golden Gate Assembly. Although Gibson and Golden Gate rely on specialized enzymes and sequence‑designed overlaps to assemble DNA fragments in a single reaction, the restriction‑digest‑plus‑ligation approach achieves the same overall goal—joining an insert into a plasmid backbone—through sequential enzymatic steps.

Figure 2. Handwritten recipe of restriction digest and ligation by Dr. Andrew Scarpelli.

During this experiment, we began by performing a plasmid miniprep to isolate the GFP‑containing plasmid backbone from E. coli, which served as the host organism carrying the engineered plasmid we intended to modify. Using the alkaline lysis method, the bacterial pellet was resuspended in Tris‑EDTA buffer to protect the DNA, lysed with NaOH and SDS to denature membranes and proteins, and then neutralized with potassium acetate to allow the plasmid DNA to renature while genomic DNA and cellular debris precipitated. After centrifugation, the plasmid‑rich supernatant was collected, and the DNA was precipitated with alcohol, washed, and resuspended to obtain purified plasmid backbone. This plasmid was then used in a restriction enzyme digest, prepared according to the reaction recipe in the notes—18 µL of plasmid DNA mixed with 2 µL of 10× buffer to create a 20 µL reaction—followed by addition of restriction enzymes to cut the plasmid at defined sites and remove the GFP region. In the subsequent ligation step, the digested plasmid backbone from E. coli was combined with the RFP insert by preparing an 18 µL ligation master mix and adding 2 µL of each DNA component to reach a 20 µL total volume, this time including T4 DNA ligase and ligase buffer to covalently join the compatible ends. Overall, the workflow followed the classic cloning pipeline: grow E. coli → isolate the plasmid backbone via miniprep → restriction digest → ligation to replace GFP with the RFP insert.

Week 7 Lab: Restriction Enzyme Digest II

Week 9 Lab: Cell Free