A Craft in the Life: Candy Cane Chocolate Bark

For a long time I have wanted to try making my own chocolate bark, as it always looks really interesting and delicious but of course is super simple! I am completely addicted to the delicious peppermint taste of candy canes and as you might expect they are pretty difficult to find when it’s not Christmas. So when I was out shopping in Lincoln and saw some I knew I had to buy them. I was thinking of making some milk and white chocolate bark with a mixture of nuts and cranberries, however after buying my candy canes I couldn’t think of anything better than milk and white chocolate bark covered in shards of smashed candy canes 🙂

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Candy Cane Chocolate Bark

You will need:

420g ‘Base’ Chocolate – Milk/Dark Chocolate (Choice of chocolate is completely up to you! If you prefer just milk, use all milk – if you would like a mixture, use half milk and half dark)

180g ‘Top’ Chocolate – White Chocolate (Again you can chose to leave this out or use white for the base of the bark and milk/dark chocolate for the top)

6 Candy Canes (I chose to smash up 5 then break 1 into larger pieces to add some more texture – but you can do whatever takes your fancy!)

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Let’s get started!

  • Begin by lining a large flat baking sheet/tray with grease proof/non stick baking paper (Ensure that your paper is larger than your tray and that the tray has a slight raised edge to prevent chocolate from spilling off the end).
  • Put your candy canes into a plastic food bag, seal and break into small pieces using a rolling pin or in my case a wooden spoon! (How much you break them up is up to you, but you can always keep a few to break into larger pieces later).

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  • Melt your chosen chocolates in separate bowls, either over simmering water (Ensuring the bottom of the bowl doesn’t touch the water) or carefully in a microwave (A lot of people say to me that they find using a microwaves burns their chocolate, however I always use this method with no problems. To prevent burnt chocolate, try removing the chocolate frequently and give it a stir. Often although the chocolate doesn’t look melted the residual heat is enough to melt all the chocolate after a good mix, and will mean you need to apply less heat overall – which should prevent burning).

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  • Leave to cool slightly, before pouring all of your ‘base’ chocolate onto the tray, spreading out evenly using the back of a spoon or palette knife.

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  • Once happy, gently pour your ‘top’ chocolate unevenly across the ‘base’ (You can either do this a bit at a time, or pour it all on at once like me!).
  • Taking a table knife or cocktail stick, drag the different chocolates into one another. Be creative! You can create swirls and even hearts (Could be good for homemade Valentine’s day treats…) with a little practise and playing around.

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  • When finished, gently sprinkle your smashed candy canes onto the chocolate, making sure to cover as much as possible.

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  • You can then add any extra larger pieces you have saved.
  • Wrap in cling film or cover in a tea towel and allow to set in the freezer for around 10 minutes.
  • Once set you can now chop or break the bark into pieces.
  • You are all done! 🙂

What is there not to love about chocolate bark? Not only does it look beautiful no matter what you put on it, but it is so quick and easy to make – you don’t even need to turn on the oven. I love how this bark turned out, the shards of candy cane give a delicious peppermint taste and bright red & white colour, while the mixture of white and milk chocolate give a marbled look that makes a great tasting base. Another great thing about chocolate bark is how easy it is to personalise, I think it would taste great sprinkled with broken pieces of honeycomb, caramelised nuts, mini marshmallows, popcorn or coffee beans!

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Your imagination and creativity can really run wild with this recipe, and you can make it for anyone and any occasion – milk/white chocolate covered in sweets for a children’s birthday party or dark chocolate sprinkled with pieces of crystallised ginger for a more grown up treat. Store your chocolate bark in the fridge, or you do risk it warming up and melting! Definitely give this recipe ago 🙂

A Craft in the Life…

A Craft in the Life: Valentine’s Day Candles

If you read my Candle Making post you will remember my candle making experience with Megan from Megan’s Candle Corner! She is currently stocking Valentine’s candles, including some very cute heart shaped tea lights that would make a great gift for anyone who loves candles 🙂

Megan's Candles

As with a lot of Megan’s candles, these tea lights are also fully customisable to suit your tastes. You can choose from scented (Forest fruits or something else from her delicious smelling collection!) or unscented if you prefer and of course your favourite colour (Pink, red, purple, turquoise, yellow, white or mixed).

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Definitely check out her Etsy and Facebook pages if you are looking for a thoughtful and handmade gift for Valentine’s day, or for any other special occasion. As she has plenty of other candles to choose from, including beautiful tea cup & saucer candles and jar candles  🙂

A Craft in the Life…

A Bake in the Life: Carrot and Walnut Cake

Since buying my new Sweet Things recipe book I have been distracted from the world of baking and focused on the art of chocolate, sweet and candy making. As a result I decided that I definitely needed to get back into baking and try out a new cake recipe that I hadn’t tried before. I am a massive fan of carrot cake, however for what ever reason I have never actually made one myself. With that in mind I decided that I really had to give it a go and to go all out with a double layered carrot and walnut cake with plenty of sweet cream cheese icing 🙂

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Carrot and Walnut Cake

You will need:

9oz/250g Self Raising Flour

2 tsp Mixed Spice/Cinnamon (I chose to use mixed spice as I already had some in my cupboard, however you can use what ever spices you like best/or have available)

9oz/250g Caster Sugar

12 fl oz/350ml Vegetable Oil

4 Eggs

12oz/350g Grated Carrots

3½oz/100g Chopped Walnuts (You can leave these out if you like or use a different nut, such as pecans)

Icing

7oz/200g Cream Cheese

4½oz/125g Softened Butter

1lb/450g Icing Sugar

1 tsp Vanilla Flavouring/Extract

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Let’s get started!

  • Preheat your oven to 180ºC/350ºF/Gas Mark 4.
  • Grease and line two 23cm round cake tins with grease proof paper or non stick paper.
  • In a large bowl, combine flour, chosen spices and sugar.

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  • In a measuring jug combine the oil and eggs.
  • Add the wet ingredients to the dry and beat well, before stirring in the carrot and chosen nuts if using.

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  • Divide the cake mixture evenly between the two tins.

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  • Bake for 25-30 minutes until light brown, firm to the touch and a skewer comes out clean from the centre.
  • Cool cakes on a cooling rack before removing.
  • To prepare the cream cheese icing, cream together the butter and cream cheese until smooth.
  • Add the icing sugar and vanilla then beat until combined.

 

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  • Once the cakes are completely cool, use the icing to fill the cake and if you like ice the top and sides.
  • I choose to sprinkle some broken pieces of walnuts on top of the cake to decorate.
  • You are all done 🙂

What a easy and simple cake! It feels as though you are using way too much carrot, however once combined and baked with everything else those worries definitely disappear. The original recipe I took inspiration from on Allrecipes used 14oz/400g of caster sugar, which seemed far too much for my taste, especially with the sweet cream cheese icing so I decided to reduce it to 9oz/250g. I also slightly adjusted the icing measurements as it produced a very large amount of icing which was far too runny – it actually may be worth only making half the amounts here to start with, then making more if you think you need it (Especially if you aren’t planning on icing the whole cake, just filling it and topping it). I really like the idea of using pecans instead of walnuts next time I make this cake, as I think it would give a really interesting change to the fairly traditional carrot and walnut combination.

I really like using oil in cakes rather than butter as it gives a more moist cake that stays that way for much longer than when using butter (Although I admit it does look slightly off putting when you are measuring it out, compared to butter! However, vegetable oil is actually healthier for you than butter, as it contains high levels of unsaturated fats which help to lower LDL/’bad’ cholesterol in the body, while butter is higher in saturated fats which reduce HDL/’good’ cholesterol). All in all a great cake that I definitely recommend trying out 🙂

A Bake in the Life…

A Craft in the Life: Sea Salted Caramel Truffles

A while back myself and a friend made some delicious Salted Caramel Chocolate Cookies, which turned out really well and were very popular with my friends. So when I saw this recipe for sea salted caramel truffles I knew I had to give it a try! I love the combination of very sweet caramel and chocolate with the savoury saltiness, so to me these are perfection.

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Sea Salted Caramel Truffles

You will need:

10oz/250g Milk/Dark Chocolate (You can either use half of each of just all of your favourite!)

2oz/50g Caster Sugar

4fl oz/125ml Double Cream

2oz/50g Light Brown Sugar

1oz/25g Butter

1 tsp Vanilla Flavouring/Extract

Large Pinch of Sea Salt Flakes (If you don’t have flakes you can always use table salt)

To Coat

7oz/ 200g Milk/Dark Chocolate (Again, you can either use half of each of just all of your favourite)

3½oz/100g Cocoa/Icing Sugar (You can coat half in cocoa and half in icing sugar, or just do all in your favourite)

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Let’s get started!

  • Begin by greasing and lining an 18cm square baking tray (Or something similar, this is a rectangular 17cm by 27cm tin and it worked perfectly – just make sure your tray isn’t too big or else you truffles will be very thin) with grease proof or non stick paper.
  • Finely chop chocolate into a large bowl and set aside.
  • In a small saucepan, combine caster sugar with 2-3 tbps of water and set over a low-medium heat to dissolve the sugar without stirring.
  • Bring to the boil and continue to cook the syrup until it becomes an amber coloured caramel (Gently shaking the pan to prevent it from sticking).

 

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  • Remove from the heat and add the double cream, light brown sugar, butter and vanilla.
  • The caramel will have hardened at this point, so return to the heat and boil. Stirring until remelted and thoroughly combined.

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  • Pour the hot caramel cream over the chopped chocolate, add the salt and stir briefly to combine.

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  • Leave for 2-3 minutes to allow the heat from the caramel cream to melt the chocolate, then beat until smooth.

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  • Pour into the prepared tin and spread level using the back of a spoon (You can also gently tap the tray on the work surface to even the mixture out).

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  • Allow to cool before chilling until firm (I decided to leave the ganache overnight in the fridge, which worked very well).
  • To coat, melt chocolate over a pan of simmering water or carefully in a microwave – allow to cool slightly (Or else when you try to coat your truffles they will begin to melt very quickly, which makes things rather tricky!).
  • Place the cocoa and/or icing sugar in separate bowls.

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  • Cut the ganache into bite sized rectangular truffles using a hot kitchen knife.

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  • You can either coat them like this or you can roll each truffle into a ball using the palms of your hands (You will need to wash your hands off after every 4/5 truffles as this gets very messy).
  • Drop 3/4 truffles into the melted chocolate, turning over with a fork to cover completely.

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  • Lift the truffles from the chocolate, allowing any excess chocolate to drip off before dropping into either the cocoa powder or icing sugar.

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  • Turn to coat on all sides before leaving to set for 4-5 minutes (This is very important, as the truffles do start to melt slightly from the heat of the melted chocolate and need this time to begin to cool down).
  • Remove carefully from the cocoa/icing sugar and place in a plastic container lined with grease proof paper.
  • You are all done! 🙂

 

I have never made caramel before but I found it really simple and you don’t even need a sugar thermometer! Similarly to my Cherry Coconut Bombs extra time is needed when it comes to rolling these into balls, tipping in melted chocolate and the cocoa/icing sugar – as it can be difficult to do without the truffles breaking apart and getting very messy! Definitely store these truffles in a cool place (I decided on just keeping them in the fridge), as they can re melt if warm.

The solid chocolate centre has a good mixture of sweet chocolate flavour with a touch of salt, however I found the cocoa powder to be a bit bitter and later decided to try re coating some in icing sugar to help reduce this bitterness slightly. I also decided to top the truffles with a little more salt, which I think worked well. I really enjoyed making these truffles, and they have gone down very well with friends! I would recommend giving them a try if you are as big a fan of the combination of savoury and sweet flavours as me 🙂

A Craft in the Life…

A Year in the Lab: PFGE Plug Production and Antibiotic Resistance Testing

Monday 20th January 2014

I actually started the week on Sunday night at 6.30pm, inoculating 3 10ml nutrient broths with a single colony from streak plates of 3 identified isolates of Escherichia coli. At 7pm these were incubated at 37ºC with agitation, and then removed at 11am after 16 hours incubation in order to obtain optimum growth/number of bacterial cells.

In order to reacquaint myself with genotypic analysis before beginning work with PFGE I started the week by running through:

  • DNA extraction – Phenol chloroform isoamylalcohol
  • Restriction digestion – NotI
  • Gel electrophoresis – Standard 110V 1.5hr run

DNA extraction is performed on an incubated bacterial sample, in order to extract and purify DNA from within individual bacterial cell nuclei before conducting a restriction digest. I began by ensuring I had a waterbath set at 60ºC, labelled eppendorfs and 95% ethanol on ice, along with all other prepared reagents and chemicals from last week.

How to conduct DNA extraction

  • Pipette 1ml of incubated sample into a 1.5ml eppendorf, centrifuge for 2 minutes at 13,000 rpm – forming a pellet at the bottom of the eppendorf.

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  • Carefully remove the supernatant and resuspend the pellet in 100μl of TE buffer and 2μg of Lysozyme (100μl TE buffer + 2μl 50mg/ml Lysozyme Stock).
  • Incubate for 30 minutes at 37ºC with agitation.
  • Remove and add 50μl 10% SDS.

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  • Incubate in a waterbath at 60ºC for 30 minutes.
  • Remove and centrifuge for 5 minutes at 13,000 rpm.
  • Remove the supernatant to a fresh eppendorf, discarding the old.
  • Add 100μl TE buffer and then, working in a fume cupboard, add 250μl phenol chloroform isoamylalcohol.

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  • Ensure the eppendorf is closed securely and mix for 1 minute (Mixture will appear white and cloudy).

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  • Centrifuge for 5 minutes at 13,000 rpm.
  • Again in the fume cupboard, carefully remove the top aqueous layer to a fresh eppendorf then repeat the phenol extraction by adding 250μl phenol chloroform isoamylalcohol.

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  • Ensure the eppendorf is closed securely and mix for 1 minute.
  • Centrifuge for 5 minutes at 13,000 rpm.
  • Again in the fume cupboard, carefully remove the top aqueous layer to a fresh eppendorf then add 250μl of chloroform.

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  • Ensure the eppendorf is closed securely and mix for 1 minute.
  • Centrifuge for 5 minutes at 13,000 rpm.
  • Repeat chloroform extraction if sample appears cloudy.
  • If not, remove the aqueous layer to a fresh eppendorf then add 650μl ice cold 95% ethanol.

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  • Incubate overnight on ice.

During the first 30 minute incubation in lysozyme I inoculated nutrient agar slopes, which I prepared last week, with isolated colonies from Brayford water sample (B2). I did this by taking a single colony from a streaked plate on a disposable loop and gently spreading it from the lowest part of the slope upwards. I inoculated 2 agar slopes per isolated organism and left them on my bench to slowly grow single colonies, from which I can work from over time.

During the second 30 minute incubation in 10% SDS I made up some more nutrient broth for inoculating isolated organisms in on further water sample collection. I prepared 30 10ml broths as before, dissolving the powder in distilled water, decanting into glass universals before autoclaving.

When I had reached the overnight incubation on ice stage, I decided to practise using the antibiotic disc plunger on my Mueller Hinton agar plates. I have used the plunger before during my undergraduate degree, however it has been a long time and I have never set it up entirely by myself. I selected 8 antibiotics from the collection which are always stored in the labs for practical classes to try out:

  • RD – Rifampicin
  • CN – Gentamicin
  • TE – Tetracycline
  • S – Streptomycin
  • AMP – Ampicillin
  • P – Penicillin
  • PB – Polymyxin B
  • VA – Vancomycin

How to conduct antibiotic disc testing

  • Begin by setting up an aseptic workspace, ensuring that labelled Mueller Hinton agar plates, samples to be spread, plastic spreaders, appropriate pipette tips and pipette are within reach.
  • Remove the lid of chosen overnight culture, flame the neck and then remove 100μl of sample, flaming the neck again before replacing the lid.
  • Lifting the lid of your labelled Mueller Hinton agar plate, gently pipette the sample onto the surface of the plate, replacing the lid and disposing of the tip.
  • Take a fresh spreader, lift the plate lid and gently spread the sample evenly over the surface of the agar.

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  • Dispose of the spreader, replace the lid and leave the plate to dry for around 10 minutes.
  • Load the antibiotic disc plunger by gently pushing your chosen antibiotic tubes into the designated holes – listening for a click.

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  • Once dry, remove the lid of your chosen plate, place the plunger over the top of the plate so it is sitting on the bench.
  • Firmly and quickly push down on the plunger to release the discs onto the agar.

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  • Remove the plunger, check all discs are present then replace the plate lid.
  • Occasionally discs may not be released, if this happens remove the missing antibiotic disc tube from the plunger and set aside.
  • Dip forceps in ethanol and then flame to sterlise, before allowing to cool.
  • Remove 1 disc from the tube and carefully place onto the agar plate.
  • Ethanol and flame the forceps again before touching another disc or returning to the bench.
  • Incubate overnight at 37ºC.

Tuesday 21st January 2014

I began my day by finishing off DNA extraction:

  • Centrifuge for 5 minutes at 13,000 rpm.
  • Remove supernatant and resuspend the pellet in 70% ethanol.
  • Centrifuge for 5 minutes at 13,000 rpm.
  • Remove and discard the supernatant, then air dry the eppendorf until all ethanol has evaporated off.
  • Add 30μl of distilled water and allow to incubate at room temperature for 5 minutes.

I then continued by conducting a restriction digest on my extracted DNA, in order to ‘cut’ the DNA at specific points, creating fragments which can be seen and measured through gel electrophoresis. Different restriction enzymes can be chosen in order to cut DNA at specific places in the genetic code. For example, the enzyme I am using – NotI cuts DNA at GC/GGCCGC and was recommended for use in a protocol I am using to run identified Escherichia coli samples on PFGE.

How to conduct a restriction digestion

  • In a 1.5ml eppendorf, combine extracted DNA with chosen restriction enzyme, appropriate 10x restriction enzyme buffer and distilled water to make up to 30μl.
  • Mix gently to combine.
  • Incubate for an hour at 37ºC.

While waiting for the restriction digestion to incubate I observed the Mueller Hinton agar plates I incubated overnight. Using a ruler I measured the zones of inhibition visible around each antibiotic disc, noting down the result and photographing each plate. This measurement can then be compared against a reference standard supplied with the discs.

Generally speaking, the presence of clearing around certain antibiotics demonstrates the effectiveness of the antibiotic against the bacteria present on the plate. If a bacterial isolate is susceptible to an antibiotic the zone of inhibition will be larger due to it’s ability to inhibit growth of bacteria. Conversely if the bacteria demonstrates resistance to an antibiotic the zone of inhibition will be smaller/not present due to the inability of the antibiotic to prevent growth of the bacteria.

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I did notice that my first batch of Mueller Hinton agar plates were too thick, so I decided to make a second batch paying special attention to the thickness of each plate as I poured aseptically.

Once restriction digest incubation had finished, I began gel electrophoresis in order to separate DNA fragments present by individual size and charge. As a result, the smaller and more negatively charged molecules move faster, while the larger and less negatively charged molecules move slower through the gel. Therefore separation occurs due to individual electrophoretic mobility, and bands produced can be compared to known sized ladders.

How to prepare a 0.8% agarose gel

  • Begin by using masking tape to seal a gel casting box, making sure there is a tight seal between tape and box.
  • Place on a flat surface away from disturbances and position the comb at one end of the gel – ensuring it doesn’t touch the bottom of the tray.
  • Prepare a 0.8% agarose gel by combining 0.8g of agarose powder with 100ml of 1x TAE buffer in a conical flask, topping with grease proof paper or a large lid to prevent fluid loss through evaporation.

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  • Heat in a microwave for around 2 minutes, using short bursts to ensure boiling and gently swirling to combine.
  • Once gel appears clear and homogeneous remove carefully and leave to cool on bench.

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  • When you can comfortably hold the base of the flask in your hand, carefully pour the cooled agarose into the taped tray, ensuring no bubbles are formed.

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  • While the solution is still liquid you can use a clean pipette tip to pop/drag any bubbles to the sides, but be careful not to do this once setting has begun.
  • Allow to set for at least 15 minutes.

How to load and run gel electrophoresis

  • Begin by gently removing the comb from the set agarose gel, removing tape around gel box and placing it into an electrophoresis tank.

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  • Ensure that your wells are nearest the negative electrode or black lead, as DNA is negatively charged and will move towards the positive electrode or red lead.
  • Produce 1L of 1x TAE buffer by combining 20ml of 50x TAE buffer stock with 980ml of distilled water in a volumetric flask and gently combine.

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  • Fill both ends of the electrophoresis tank, allowing the buffer to cover the agarose gel by around 3mm, but not reach the overfill line.
  • Prepare loading solutions in 1.5ml eppendorfs before carefully pipetting 20μl of each sample into a separate well, including a loading buffer to allow visualisation of DNA fragment progression (This does take a bit of practise as you need a steady hand and to release the solution just above the well, removing the pipette from the buffer before releasing entirely).

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  • Pipette your chosen appropriate ladder to allow comparison and calculation of fragement/band size.
  • Ensure the power pack is switched off, place the lid onto the electrophoresis tank.

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  • Attach the leads to the power pack, set to your requirements (In my case 110V for 1.5 hours) and confirm.
  • Check that bubbles are forming at both ends of the tank then leave to run.

How to stain and view the gel

  • Once finished, switch off the power pack and remove the lid.
  • Wearing gloves, gently lift the gel box out of the tank and slide the gel off into a large plastic tray.
  • Tip the TAE buffer into the tray and add 5μl Ethidium Bromide to the buffer (As Ethidium Bromide is carcinogenic ensure that you are wearing gloves and dispose of used equipment appropriately).

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  • Allow to stain for 30 minutes before visulising using an imaging system – in my case Kodak Gel Logic 100.
  • When finished dispose of your gel into the same carcinogenic waste as before.

Once finished, gel observed and the image saved I reviewed and prepared PFGE reagents and chemicals for use tomorrow. I decided to repeat antibiotic testing using my new thinner Mueller Hinton agar plates, following the same procedure as yesterday – incubating overnight at 37ºC.

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In preparation for plugs production tomorrow I inoculated 3 10ml nutrient broths with an individual single colony of 3 identified isolates of Escherichia coli. At 7pm these incubated at 37ºC with agitation, and were removed at 11am after 16 hours incubation in order to obtain optimum growth/number of bacterial cells.

Wednesday 22nd January 2014

I started the day by removing my overnight nutrient broth sample and beginning PFGE plug preparation.

How to produce PFGE bacterial plugs

  • Add chloramphenicol and incubate for an hour at 37ºC with agitation.

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  • Melt the 2% agarose for a few seconds in the microwave before equilibrating the solution at 50ºC in a waterbath.
  • Centrifuge all 10ml of each sample in separate eppendorfs, in 1.5ml batches for 3 minutes at 130,000 rpm, removing and discarding the supernatant each time.
  • Resuspend the pellet in cell suspension buffer and equilibrate at 50ºC in a waterbath (I decided to make 1ml of 1% plugs and therefore the amount of cell suspension buffer and 2% agarose were the same – 500μl).

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  • Combine 2% agarose with cell suspension and mix gently.
  • Carefully pipette into plastic molds (85μl) and allow to solidify before moving to 4ºC for 10-15 minutes to add strength to the plugs before removal.

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While samples were undergoing chloramphenicol incubation I observed my antibiotic resistance testing plates, measuring zones of inhibition, making notes and taking photographs.

During plug pipetting it became very clear that the cell concentration within the plugs was too high, and even though plugs were left to set for an hour the integrity was too fragile to continue. I decided to try a lower initial amount of sample and 3 more 10ml nutrient broths were inoculated with an individual single colony of 3 identified isolates of Escherichia coli. Again these were incubated at 37ºC with agitation for 16 hours.

Thursday 23rd January 2014

Again, I began my day by removing my overnight nutrient broth sample and beginning PFGE plug preparation. I conducted the production in the same way as previously, however I decided to just use 2ml of sample. As I reached the pipetting stage the plugs felt a lot more stable and on removing them into individual eppendorfs they retained their integrity very well. I therefore continued the process:

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  • Remove each plug into an individual eppendorf.
  • Add lysozyme to each plug and gently mix.
  • Incubate for 2 hours at 37ºC.
  • Remove each plug gently from lysozyme, rinse with distilled water and replace into individual eppendorfs.

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  • Add proteinase K to each plug and gently mix.
  • Incubate overnight at 50ºC.

Once finished I decided to conduct some more antibiotic resistant testing, using 3 identified Escherichia coli samples from the bacterial isolate collection. I used the same procedure as previously, including the same antibiotics and decided to conduct repeats of each. Incubating overnight at 37ºC.

Friday 24th January 2014

I had a supervisor meeting from 10am-12pm, however as I needed to conduct the next washing stage of my plug production I began work at just before 10am. I started by preparing 1x wash buffer from a 10x wash buffer stock.

PFGE plug production: Washing Stage

  • Wash each plug in 1ml of 1x wash buffer for an hour each, with agitation at room temperature.
  • Repeat 4 times.
  • Store in 1x wash buffer at 4ºC.

Once completed I observed my overnight antibiotic resistance plates, recorded the zones of inhibition present and photographed each plate.

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Next week…

Now I have a first batch of plugs prepared I can conduct a restriction digest on the plugs on Monday, incubating overnight before running on Tuesday. Once the results from these plugs have been visualised I can either adjust my procedure to improve them or begin repeating the procedure on other identified Escherichia coli samples from the bacterial isolate collection. I am very happy with the antibiotic resistance testing procedure I have conducted this week, so I will continue researching, ordering and preparing reagents and chemicals for IEF next week.

After my supervisor meeting on Friday some aspects of my project have changed slightly so I will also be adjusting my research plans for the coming months and filling in a GS4 next week. Hopefully beginning my first large scale on site sample collection in the first week of February 2014!

A Year in the Lab…