Working with RNA & DNA

Question 1

what is whole mount in situ-hybridisation?

Answer 1

visualises the spatial distribution/location of mRNA expression within an intact, whole organism or tissue

Question 2

list the three controls that should be used in a WMISH initial experiment testing expression of a known gene & why

Answer 2

antisense RNA probe
- positive control - binds to complementary mRNA of target gene & should produce a signal
sense RNA probe
- negative control - detects non-specific binding or background staining; should produce NO signal
knockout embryo
- embryo lacking gene of interest; should show no signal even with antisense probe present as target mRNA isn’t present

Question 3

list three conditions should be included in a WMISH analysing the expression pattern of a novel gene, to ensure both a positive and negative control - and why?

Answer 3

1. experimental antisense RNA probe (positive control) - antisense riboprobe to novel gene, detects expression patter
2. sense RNA probe (negative control) - sense riboprobe to novel gene should show no signal - rules out non-specific binding
3. antisense RNA probe for known gene (positive control) - shows that reagents & protocols are working as expected

Question 4

difference between antisense and sense probes?

Answer 4

antisense probe = complementary to target mRNA; binds specifically to it, detects gene expression & gives signal

sense probe = negative control; same sequence as mRNA, doesn’t bind to target mRNA but checks for background staining & nonispecific binding

Question 5

steps of WMISH?

Answer 5

1. FIXATION= embryos/tissues fixed with formaldehyde
   - preserves tissue structure & prevent RNA degradation
2. WASHING = after fixing, formaldehyde is washed off/removed with several washes of PBS + Tween
   - removes fixatives and permeabilize tissue

3.. DEHYDRATION = embryos are dehydrated through a methanol series (25-50-75-100% methanol) - stored at 100% methanol
- removes lipids, increases probe penetration, and stores embryos long-term

4. REHYDRATION = return embryos to aqueous conditions for probe application through gradual methanol dilutions (100-25%- PBS)
5. PERMEABILISATION = proteinase K digestion especially for older embryos
   - breaks down proteins to help riboprobe diffuse into tissues
6. HYBRIDISATION WITH RIBOPROBE = add labelled antisense-RNA probe complementary to target mRNA; incubate sample mRNA with probe in hybridisation buffer at approx. 65 degrees
7. POST-HYBRIDISATION WASHES = RNAse treatment (RNAse A & T1 digest) unbound/ single-stranded RNA
   - removes non-specific or weak riboprobe-mRNA binding
8. DETECTION
   - riboprobe has an incorporated label (e.g. DIG - hapten-tag)
   - use anti-DIG antibody conjugated with an enzyme (e.g. alkaline phosphatase/ horseradish peroxidase)
   - then add substrate into mix (e.g. AP substrate is BCIP + NBT; horseradish peroxidase + DAB substrate)
9. MOUNTING & IMAGING = store in 70% glycerol for long-term storage and imaging
   - mount on slides, observe under light microscope

Question 6

describe the process of using a DIG-labelled riboprobe in ISH - how would you detect a signal?

Answer 6

1. RNA probe is made in vitro from a DNA template using RNA polymerase - one NTP (usually UTP/uracil) has a DIG label attached
2. DIG-labelled NTP in riboprobe undergoes complementary binding to target mRNA region
3. anti-DIG antibody conjugated with enzyme (e.g. AP/HRP) binds to DIG tag
4. substrate to enzyme added:
   - AP + BCIP & NBT substrate = produces blue/purple precipitate
   - HRP + DAB substrate = produces brown signal or fluorescent signal
5. signals visualised under light microscope - can analyse expression pattern in sample

Question 7

what is formaldehyde? why is it used to fix tissues and embryos for ISH?

Answer 7

formaldehyde - a cross-linking agent that stabilises proteins and protects against RNases
- preserves tissue structure & prevent RNA degradation

Question 8

what is used to remove the formaldehyde from the embryo?

Answer 8

PBS + Tween - removes fixative & permeabilises tissue

Question 9

You perform WMISH using a labelled antisense probe for a gene of interest, but observe no signal.
What two possible explanations could account for this? (2 marks)

Answer 9

1. antisense riboprobe isn’t complementary to the target mRNA of the gene of interest - no binding = no signal
2. gene of interest isn’t expressed at that stage or in the sample itself
   3, detection system failed to detect gene expression

Question 10

Why is it important to include a sense probe control in a WMISH experiment? (3 marks)

Answer 10

sense probe is a negative control - sense probe has the same sequence as the target mRNA
- as sense probe is non-complementary = shouldn’t hybridise = no signal expected
- ensures there’s no non-specific binding or background staining that could affect data
- confirms that any signal from antisense probe is specific to mRNA

Question 11

During WMISH, you forgot to perform the proteinase K digestion step.
What effect might this have on the result, and why? (4 marks)

Answer 11

proteinase K digests proteins and increases tissue permeability - especially if it’s an older embryo (e.g. >24hpf zebrafish embryo)
- no proteinase K = lack of permeabilization = harder for riboprobe to enter sample & bind to complementary target mRNA of gene of interest = harder to visualise gene expression patterns in embryo
- results in weak/ no hybridisation signal

Question 12

You observe weak background staining across your whole embryo, even in the sense probe control.
Give three potential sources of non-specific signal. (3 marks)

Answer 12

• incomplete washing of unbound/weakly-bound riboprobes
• over digestion by proteinase K/ poor tissue integrity
• probe cross-hybridising with similar but not-target mRNA sequences

Question 13

Design a WMISH experiment to test whether a gene is expressed in the neural tube of zebrafish embryos.
Include controls and justify each step briefly. (6 marks)

Answer 13

1. fix embryos with formaldehyde to preserve structure and RNA (protect against RNases)
2. washing to remove formaldehyde and permeabilise tissue using PBS + Tween
3. permeabilise embryo with proteinase K - digests proteins, increases tissue permeability - allows probe entry
4. (negative control step) use sense probe first to check for non-specific binding - no signal = no non-specific binding or background staining
5. use antisense riboprobe (complementary to target mRNA of neural tube gene) as a positive control
6. riboprobe conjugated with tag (e.g. DIG tag)
   - add anti-DIG antibody conjugated with enzyme (e.g. alkaline phosphatase)
   - add substrate to AP (NBT + BCIP)
   - detect blue/purple precipitate from enzyme-substrate reaction under light microscope

Question 14

Explain the importance of the RNase treatment step following hybridisation in WMISH.
What would happen if this step was skipped? ( 5 marks)

Answer 14

RNAse treatment contains RNAse A and T1 - removes any unbound or weakly-bound riboprobes from the solution following hybridisation through a series of washes
- leaves only complementary riboprobe-mRNA binding = improves specificity of signal & gene expression pattern data

what would happen - unbound/ weakly bound riboprobes would remain
- high background signal/ inaccurate data
- misinterpret expression pattern

Question 15

A student accidentally uses the sense probe on all their embryos instead of the antisense probe.
They still observe a signal.
What does this suggest about the result, and what should they do next? (4 marks)

Answer 15

• sense probe has the same sequence as the target mRNA of the GOI - there shouldn’t be a signal
  -presence of a signal suggests:
• non-specific binding
• high background staining
• sense probe was synthesised incorrectly
• faulty reagents
• previous antisense result may be a false positive

what to do:
- repeat with a properly synthesised antisense probe for GOI (as a positive control) = confirms correct orientation, function, and signal for probe. this also checks reagents & hybridisation conditions are working
- resynthesizing the sense probe = potentially incorrect synthesis initially
- adding KO embryos & then adding probes (sense and antisense) = confirms probe specificity (should be no signal) and procedure, reagents…
- check embryo prep quality = e.g. proteinase K step must be done properly for permeabilising tissue for riboprobe activity

Question 16

Why is methanol dehydration used during the WMISH procedure? (3 marks)

Answer 16

gradual methanol from 25-50-75-100% methanol for dehydration
- sample can be stored in 100% methanol
- removes lipids from sample
- increases riboprobe penetration
- helps preserve tissue morphology & allows for long-time storage of embryos

Question 17

You run a WMISH using a probe to a housekeeping gene, and observe no expression.
You also see no signal with your gene of interest.
What conclusions can you make about the experiment, and what steps should you take? (5 marks)

Answer 17

house-keeping genes are consistently active genes in all cells so there should be a signal - act as internal positive controls
- something wrong with protocol - e.g. riboprobe prep, reagents, embryo prep, faulty detection
- can’t conclude anything about novel gene expression
- need to repeat with a fresh probe, checking fixation, digestion & riboprobe synthesis steps
- validate hybridisation conditions and antibody detection system

Question 18

what feature allows for the riboprobe to be detected within a tissue?

Answer 18

riboprobe detected as it’s conjugated with a detectable marker
- fluorescent tag (GFP)
- biotin = reacts with streptavidin, conjugated with enzyme - undergoes enzyme-substrate reaction = produces colorimetric/measurable signal
- enzyme-conjugated tag

Question 19

what equipment is needed for a general reverse transcription reaction? (6)

Answer 19

reverse transcription - synthesises cDNA strand from mRNA
- mRNA template
- reverse transcriptase
- reaction buffer = Mg2+ ions, salts…
- dNTPs
- RNAse inhibitor = prevents RNA degradation
- gene-specific primer

Question 20

What are the key reagents used in riboprobe synthesis and why are they needed?

Answer 20

DNA Template: Contains the gene of interest for transcription.

RNA Polymerase (T7, T3, SP6): Synthesizes RNA from the DNA template.

Nucleotide Mix (ATP, GTP, CTP, UTP): Provides the building blocks for RNA synthesis.

Labeling Nucleotide (e.g., DIG-UTP, Biotin-UTP): Labels the probe for detection.

Transcription Buffer: Ensures optimal ionic conditions for RNA polymerase activity.

RNase-free Water: Prevents RNA degradation during the reaction.

Question 21

What controls are needed in riboprobe synthesis and why?

Answer 21

No Template Control: Ensures there’s no contamination or non-specific transcription.

Template Control: Confirms that the RNA polymerase is functioning and the template is viable.

RNase Control: Verifies that no RNA degradation occurs during synthesis.

Question 22

What is the purpose of synthesizing a riboprobe?

Answer 22

to generate a labelled RNA probe that can detect specific RNA or DNA sequences in hybridization assays or localization studies

Question 23

protocol for riboprobe synthesis for ISH (complementary to gene of interest/GOI)

Answer 23

1. isolate mRNA from sample expressing GOI (using poly-T columns/ beads)
2. reverse transcription reaction (synthesise cDNA from mRNA template using reverse transcriptase)
3. amplify cDNA using PCR (primers specific to cDNA, Taq polymerase)
4. clone PCR product into a plasmid via T-A cloning (Taq polymerase leaves A overhangs, plasmid has T overhangs).
5. in vitro transcription: Use T7/ SP6 RNA polymerase to transcribe the cloned cDNA into an RNA riboprobe.

Question 24

What is Hyb+ used for in zebrafish in situ hybridization (ISH), and what does it contain?

Answer 24

Hyb+ is a hybridization buffer used during pre-hybridization and hybridization steps in zebrafish ISH.

Purpose:
- Reduces non-specific binding
- Stabilizes probe-mRNA hybridization
- Enhances probe penetration

Contains:
- Formamide (lowers Tm)
- SSC, tRNA/heparin (block background)
- Tween-20 (permeabilizes tissue)

Question 25

what is the purpose of PCR?

Answer 25

amplifies a specific DNA sequence from cDNA (or genomic DNA) using primers and Taq polymerase in a thermal cycler

Question 26

what is required for a PCR reaction?

Answer 26

- cDNA (template for amplification) - forward & reverse primers (starting points for DNA synthesis) - Taq polymerase (heat stable enzyme; adds nucleotides) - dNTPs - buffer (maintains optimal pH/salts)

Question 27

steps in PCR?

Answer 27

1. **denaturation (95 degrees)** - breaks hydrogen binds between complementary base pairs in ds DNA - melts dsDNA to ssDNA strands 2. **annealing (50-65 degrees)** - forward & reverse primers anneal to their complementary sequences on ssDNA - forward primer binds to one template strand; reverse primer binds to the opposite strand at the opposite ends 3. extension (72 degrees) - Taq polymerase adds dNTPs to the 3’ end of each primer, synthesising new DNA strands - now have two copies of your DNA region of interest – one from each strand

Question 28

why is denaturation in PCR reactions conducted at 95 degrees?

Answer 28

high temperature disrupts base-pairing = breaks H-bonds = melts DNA duplex/dsDNA

Question 29

why is the temperature for annealing in PCR between 50-65 degrees? why is it so variable?

Answer 29

annealing temperature allows forward & reverse primers to anneal to complementary DNA sequences on ssDNA variable as temperature depends on the melting temperature (Tm) of the primers - must be low enough for H-bonds between primer & strand to form, but high enough to prevent non-specific binding

Question 30

what is the significance of the forward & reverse primers in PCR?

Answer 30

forward primer binds/anneals to complementary sequence on one template DNA strand reverse primer binds to complementary sequence on opposite end of opposite/other strand mark start and end points for DNA synthesis & amplification in 3' direction

Question 31

why is the temperature for extension in PCR at 72 degrees?

Answer 31

optimal working temperature range for Taq polymerase (head-stable enzyme) - adds dNTPs to 3' end of primer (synthesis in 5-3' direction)

Question 32

Why is gel electrophoresis used after PCR?

Answer 32

checks the presence, size & purity of the amplified DNA fragments

Question 33

what is the importance of DNA purification after PCR?

Answer 33

removes leftover primers, enzymes, salts, and dNTPs - get clean PCR product for downstream experiments (e.g. cloning, sequencing)

Question 34

How does DNA move through a gel in electrophoresis?

Answer 34

DNA is negatively charged - migrates towards the positive electrode through the agarose gel - smaller fragments travel further and faster

Question 35

What is the function of a DNA ladder in gel electrophoresis?

Answer 35

acts as a molecular weight marker to estimate the size of DNA fragments in the sample

Question 36

how is ethidium bromide used as a dye to stain DNA following gel electrophoresis for visualisation?

Answer 36

ethidium bromide - intercalates between DNA bases and fluoresces under UV light (blue)

Question 37

why are GelRed or SYBR Safe better alternatives to ethidium bromide for visualising DNA fragments following gel electrophoresis?

Answer 37

less toxic work with UV/ blue light

Question 38

what is the use of loading dye in gel electrophoresis?

Answer 38

loading dye weighs down the DNA so it sinks into the well

Question 39

troubleshooting in gel electrophoresis - no DNA bands visible, but DNA ladder is present. What might have gone wrong?

Answer 39

- failed PCR reaction - no DNA present in same or sample degraded - DNA didn't mix with loading dye

Question 40

troubleshooting in gel electrophoresis - DNA ladder is missing but sample bands are present. What’s the likely error? How can we prevent this happening?

Answer 40

errors: - forgot to load the DNA ladder sample - poor mixing with loading dye = ladder didn't sink properly improvements: - double-check ladder has been added & to the correct well - mix ladder well with loading dye and briefly centrifuge if needed

Question 41

In a visualisation of a gel electrophoresis of DNA following a PCR reaction, the DNA bands appeared smeared. What could be the cause? How can this be prevented?

Answer 41

errors: - overloading wells with DNA sample - degraded DNA (DNA broken down before loading) - poor reagents or buffer contamination improvements: - use fresh DNA, keep samples on ice, add RNase if needed to prevent degradation - use less DNA (don't overwhelm wells) - use good quality agarose & fresh buffers/reagents

Question 42

In a visualisation of a gel electrophoresis of DNA following a PCR reaction - all bands appeared faint. What could have gone wrong? How can this be prevented?

Answer 42

errors: - not enough DNA loaded into wells/ DNA is too low concentration - UV/blue light exposure is weak - insufficient staining for visualisation improvements: - use more template, concentrate your sample, or increase PCR yield = prevents having low/insufficient DNA concentration - make sure gel or sample contains enough stain (e.g. GelRed) - adjust imaging settings/use a stain more sensitive to UV or blue light

Question 43

In a visualisation of a gel electrophoresis of DNA following a PCR reaction - bands appear in the wrong location (unexpected sizes). What might this indicate?

Answer 43

errors: - incorrect DNA ladder = size estimation is wrong - small faint bands near the bottom of the gel may mean primers bound to each other (primer-dimers), not the sample; or template contamination with other cDNA - multiple bands or at the wrong sizes may mean there were non-specific PCR products produced improvements - double-check ladder type matches target bp range - prevent non-specific PCR products by adjusting annealing temp, redesign primers, reduce cycle number - prevent primer-dimers = raise annealing temp to increase specificity, reduce non-specific binding with hot-start Taq polymerase - prevent template contamination with other cDNA by using clean techniques

Question 44

what are the two ways in which protein synthesis is regulated?

Answer 44

1) level of the transcript - eukaryotic mRNA has structural features that affect translation efficiency - e.g. complex secondary structures in the 5' UTR can hinder ribosome scanning and reduce initiation at the start codon 2) global/cell-wide regulation - under stress conditions, cells broadly inhibit protein synthesis - prolonged stress may activate pathways like apoptosis to maintain cellular integrity

Question 45

main site of protein synthesis?

Answer 45

80s ribosome - made up of 40 and 60s subunits that associate & disassociate - maximal translation potential occurs when they're disassociated - 60s subunit contains peptidyl transferase = links amino acids via peptide bonds

Question 46

what is the importance in using different detergents (e.g. SDS vs Triton X-100) to extract proteins from different cellular compartments (e.g. nucleus vs cytoplasm)?

Answer 46

SDS - strong detergent; breaks both cytoplasmic and nuclear membranes - gives whole-cell extract (proteins from all compartments) Triton X-100 - gentler detergent; only breaks outer membrane & gives cytoplasmic protein extract

Question 47

(3 marks) Outline how an antisense RNA probe is generated from cDNA and how it is used in WMISH.

Answer 47

Clone gene of interest into a plasmid with a promoter (T7/SP6). Linearize the plasmid downstream of the insert. Perform in vitro transcription using T7/SP6 RNA polymerase, incorporating DIG-labelled UTP during riboprobe synthesis to generate a DIG-tagged antisense probe for detection in WMISH.

Question 48

(2 marks) Explain the difference between “sense” and “antisense” RNA probes, and their respective purposes in an ISH experiment.

Answer 48

Sense probe: (negative control) same sequence as target mRNA; doesn’t bind—used as a negative control to check for non-specific staining. Antisense probe: (experimental probe) complementary to mRNA; binds specifically—used to detect gene expression.

Question 49

(2.5 marks) Outline the key steps required to generate double-stranded cDNA from a tissue sample.

Answer 49

RNA extraction: Isolate total RNA from the tissue sample, often using TRIzol or column-based methods. First-strand synthesis: Use reverse transcriptase with a primer (e.g. oligo-dT or random hexamers) to synthesize the first strand of cDNA from the mRNA template. Second-strand synthesis: Use DNA polymerase to synthesize the second DNA strand, forming double-stranded cDNA.

Question 50

(3 marks) Describe the principle behind T-A cloning, including the roles of sticky ends and Taq polymerase.

Answer 50

Taq polymerase adds a single A overhang to PCR products due to its lack of proofreading activity. The plasmid vector is cut by restriction enzymes, then prepared with a complementary T overhang. The A and T overhangs anneal, and DNA ligase seals the nicks, enabling ligation of the insert into the vector.

Question 51

(2 marks) Give two advantages of using sticky-end ligation over blunt-end ligation in cloning.

Answer 51

Sticky ends can form hydrogen bonds via base pairing, making ligation more efficient than blunt ends. Sticky-end ligation can be directional, ensuring the insert goes in the correct orientation.