Plasmid Mapping

you will determine the orientation of the insert in the plasmid provided to you

  • picking the colony, growing the liquid culture and the miniprep to isolate the plasmid have been done for you

  • because this has been done for you, you may not have plasmids from the same liquid culture as the spent culture you used/will use to obtain your standard curve

Plasmid Mapping OLM

depending on how RD/ligation was performed, insert can be ligated in 2 different orientations

  • right way or upside down

using just 1 restriction enzyme → restriction digest sites on either side of the insert are identical

  • can be ligated in 2 different ways: 5’ to 3’ or 3’ to 5’

using 2 different enzymes to digest then ligate insert in

  • different restriction enzyme cut sites; sticky ends don’t match

want to express insert — must be in correct orientation

must be in the 5’ to 3’ orientation

only used one restriction enzyme, EcoR1 → insert can actually go in either 5’ to 3’ or 3’ to 5’

  • which of the plasmids have insert in correct orientation

provided with plasmid that has been isolated from miniprep made from liquid culture of picked white colony

we did not keep the same liquid cultures from spectrophotometry to plasmid mapping experiment

  • don’t necessarily match

  • if results don’t match, this is why!!!!!!!!!!!!!!

plasmid we’re using is pAMP

determine orientation of insert by looking at results of double digest

HindIII site — will cut plasmid backbone once

  • determine orientation with BamHI site

  • BamHI site is in the insert and it’s off centre of insert (towards one end)

small fragment if 3’ to 5’ and a very large segment

if 5’ to 3’: will have two fragments that are more similar in size

distinguish between orientations of insert using the pattern of the cut — i.e. the size

because we are on an agarose gel, size estimates are not super precise

  • approximate — can be off a bit (larger fragment with smaller fragment)

    • should be close

    • just looking for pattern of bands that are really close together

in lab, will be performing four different digests:

  1. uncut plasmid

  2. HindIII

  3. BamHI

  4. HindIII + BamHI

EcoRI = HindIII

combine digests (once complete) with loading dye

combine with 0.75% agarose gel and run for 30-45 min at 100V

on the gel, there is a Ladder to tell what the size of the fragments are (1 Kb Plus Ladder)

stained with Red Safe → allow us to visualize the DNA

0.75% Agarose Gel:

  • agarose is a sugar that creates pores when in gel form

  • higher the percentage, the smaller the pores

    • which means the greater the resolution (can distinguish a smaller difference in the band size)

    • however: it takes longer to run

  • maximize percentage for size of band

    • very large fragments, can get away with a lower percentage gel

  • DNA (negative) will be separated based on size when current is run → run towards positive electrode

    • separated based on size

    • smallest will run further than largest

Loading Dye:

  • must add Loading Dye to samples!

  • contains a dense sugar like glycerol

    • makes sample sink to bottom of well

  • if didn’t have a dense sugar to make it more viscous, when add sample to wells (which are sitting in running buffer), they would just float away into running buffer

  • contain a Dye (often bromothymol blue)

    • allows us to see where samples are loaded + as gel is running, allows us to visualize how far the gel has run

    • roughly 500-300 bp on agarose gel (of our percentage)

  • EDTA — stops enzymatic reactions by binding metal ions that are often required by the enzyme

    • don’t want restriction digests to keep going (nonspecific cutting)

    • PCR reaction to stop

Red Safe:

  • allows for the visualization of DNA

  • intercalates between DNA base pairs (inserts itself in between)

  • when exposed to UV, emits light and can be captured on a gel doc

    • only where there is dsDNA, we will see the light being given off → gives bands on gel

gel tank: agarose gel + running buffer (to have electricity be able to move through tank)

  • black in back! DNA is going to go closest toward black (back end) → negative

  • runs toward red (red ahead)

power pack supplies power to gel

not same gel tank or power pack we’re using but principles are the same

visualize DNA in gel dock

Gel doc: connected to computer → inside closed container, there is a camera that can sense the light (shows us the image on the computer)

  • drawer

  • place drawer

  • drawer gets closed

that image gets uploaded to OWL

What will we see?

uncut plasmid:

  • exists in multiple forms (all the sample plasmid but will run differently on the gel because of their conformations)

    • supercoiled (like a circular rubber band that has been twisted up into lots of tertiary structure) — fastest running form of plasmid

      • all condensed: will travel the fastest

    • nicked (one of the strands gets nicked but other is intact—loses supercoiled structure + runs the slowest)

      • gets hung up more

      • often occurs because of freezing and thawing

    • linear (both strands have been cut) — expect for digests (can also happen by chance)

      • middle band → lowest frequency (fainter bands)

      • difficult to be linearized without enzyme

      • amount of band = darkness

  • may form concatamers so be larger than expected

    • often get concatamers of different forms (several clumped together)

    • bigger than we expect

    • pattern of 3 different bands in uncut plasmid, representing nicked, linear, supercoiled

single and double digests:

  • single — plasmid cut once so now linear

    • cut through both strands — represented as appropriate size for plasmid + insert together

    • one singular band

  • double — plasmid cut twice so 2 binds

    • THIS IS THE DIGEST YOU USE TO DETERMINE ORIENTATION

Why Undigested and Single Digests?

  • both are controls!

  • if a lane of digested plasmid has extra bands that are the same size as undigested, shows not all the plasmid was digested

  • if double digest has extra band that are the same size as single digest, who that some of the plasmid was not digested by both enzymes

    • same size as single digest

    • some of plasmid was only digested with one of the enzymes (not both)

    • include both enzymes in single to show that both are actually working

      • explain away extra bands (if didn’t match up with other lanes, might have problem and can’t analyze gel)

  • BOTTOM LINE: IT ALLOWS YOU TO TROUBLE SHOOT ANY EXTRA BANDS THAT APPEAR

    • in either single or double digest lane

  • enzymes are often not 100% efficient — not all the plasmid is going to be cut by the restriction enzyme

    • if we have extra bands we can tell if it is what we expected by including controls

    • if a lane of digested plasmid has extra bands in it and when you look across gel, you can see that they’re the same size in undigested lane, shows that you have some that are intact and not fully digested

gel pictures will be uploaded to OWL

  • label each lane

  • identify size of bands using ladder

  • use band sizes to determine the orientation of the insert for YOUR sample

  • this will be reported in your lab report

    • tie this info back to the level of insert product

    • how would the orientation that you found for this particular sample effect the insert product level?

      • whether insert is in correct or incorrect orientation — what does that mean for production of insulin in spectrophotometry lab

How to Load a Gel

  • using a mini-sub cell to perform agarose gel electrophoresis

  • provides electric current to pass which separates the DNA fragments in the cell

  • DNA at black electrode to positive, red electrode

  • place agarose gel into chamber

  • add electrophoresis running buffers to each end

    • keep adding until covered by at least 2 mm of buffer

  • when loading samples, keep tip perpendicular to wells

  • just inside or just above well (break surface of buffer)

  • avoid bumping or movement of gel chamber

  • should see samples migrate