As they have 500ml Erlenmayer flasks here, of which a quater volume (125ml) can be used for the liquid culture, I need to use two of them, which will bring me to 250ml of liquid culture. To create the same amount again based on liquid cultures, I’ll need 2ml of liquid culture to mix with fresh LB medium.

For a 125ml of liquid culture, I’ll need:

• 125ml LB medium
• 6250μg = 6.25mg of ampicillin (a final concentration of 50μg/ml)
• some bacteria from the streaked out plate

I’ll make two of these.

I’ll also make a smaller, 100ml Erlenmayer flask control with arabinose – to see if it glows as expected. This will contain:

• 25ml LB medium
• 1250μg = 1.25mg of ampicillin (a final concentration of 50μg/ml)
• some bacteria from the streaked out plate
• 125mg or arabinose (a final concentration of 5mg/ml)

getting a 16×16 bitmap into our dropdrawer program, in this case the ChickenRunner’s icon from the age-old computer game Berzerk

letting the fab@home fabber drop arabinose on the plate

wait overnight, and see the results

The experiments done from yesterday to today show a clear result that the size of arabinose drops really matter, and that the correspondence between the brightness of the result and the size of the drop is not linear. It seems that a drop size of at least 0.02 is preferable (say a pushout of 0.03 and a suckback of 0.01), sometimes 0.015 works as well. It’s also visible that we can’t get better than a distance of 5mm between the drops, but 6mm seem to work out best. Shorter distances get along better with smaller drop sizes, probably because the liquid flows together. Bigger drop sizes also help in the process as they stick to the agar gel easier.

The following prints resulted in visible output (spacing / drop size / suckback):

• 5 / 0.03 / 0.01
• 5 / 0.03 / 0.015
• 5 / 0.025 / 0.005
• 5 / 0.02 / 0.005
• 6 / 0.025 / 0.005
• 6 / 0.02 / 0.005
• 7 / 0.02 / 0.005 (barely visible)

Whereas the following prints resulted in practically invisible results (spacing / drop size / suckback):

• 5 / 0.03 / 0.02
• 6 / 0.02 / 0.01
• 6 / 0.03 / 0.015
• 6 / 0.03 / 0.02
• 7 / 0.02 / 0.01

The following image shows three prints of spacing 6, with drops sizes below (spacing / drop size / suckback):

• 6 / 0.025 / 0.005
• 6 / 0.02 / 0.005
• 6 / 0.02 / 0.01

The following images shows prints of spacing 5, with drops sizes below (spacing / drop size / suckback):

• 5 / 0.03 / 0.01
• 5 / 0.03 / 0.015
• 5 / 0.025 / 0.005

The following image shows three different prints of different sizes (7, 6 and 5) and similar drop sizes, see below (spacing / drop size / suckback):

• 7 / 0.02 / 0.005
• 6 / 0.02 / 0.005
• 5 / 0.025 / 0.005

I’ve made a number of prints today, basically experimenting with different spacings between the drops. All images were 16×16 pixels, with the following parameters:

• spacing: 5, drop: 0.005, suckback: 0.002
• spacing: 7.5, drop: 0.005, suckback: 0.002
• spacing: 5, drop: 0.01, suckback: 0.005
• spacing: 5, drop: 0.015, suckback: 0.01
• spacing: 6, drop: 0.015, suckback: 0.01
• spacing: 7, drop: 0.015, suckback: 0.01
• spacing: 7, drop: 0.02, suckback: 0.01
• spacing: 7, drop: 0.02, suckback: 0.015

we’ll see tomorrow how they turn out. Unfortunately leveling is really an issue, especially with small drops: the syringe touches the surface of the agar on some parts of the plate, but not on the others. So drops are not placed everywhere where they should, and in some places, accumulated drops are made.

Yesterday I made a number of images to try out different resolutions and drop distances, to see which work the best. One limitation is the size of the droplet that can be reliably dropped by the fab@home 3D printer. Another is that the arabinose dropped will dissipate through the agar, in effect creating a much larger pixel than the drop itself. Also, the bacteria can overdose on the arabinose, in the end not displaying anything.

I experimented with a simple X pattern, and also with some space invaders:

The initial resolution was 16×16 pixels, with various drop sizes. The drop sizes are defined by the amount of downwards movement that is made by the syringe tool, pushing the liquid out. The spacing between the drops can also be played with, and thus I did.

The three invaders below all have 5mm between the drops, and the drop sizes are also the same, 0.03. The difference is only in the amount of suckback after a drop is made – that is, some liquid is pushed out, and then there’s a backwards movement in the syringe, sucking excess material back. The invader on the left on both images has a suckback of 0.01, while the ones on the right have none and 0.015, respectively.

It seems that no suckback resulted in an overdose of arabinose (see that the silhouette of the invader is visible, but the inside is dark). It also seems that a suckback of 0.015 was too much, though this is unclear why.

I also experimented with 32×32 pixel resolution images, with shorter distances between the drops, and smaller drops still. Here, the drops really got together during printing (see yesterday’s post), thus the result is really just blog blobs, with arabinose overdose in the middle:

The invaders, from left to right, were made with the following settings:

• spacing 3, drop 0.02, suckback 0.01
• spacing 2, drop 0.015, suckback 0.01
• spacing 1.5, drop 0.005, suckback 0.002

The next experiments will involve more spacing and smaller drops, so as th prevent arabinose overdose, and also to enhance the effect of single pixels.

It’s also a challenge to make these photos, in low light conditions it’s very difficult to focus properly. Still, thanks to Dániel Molnár for his tripod, as my own is still missing

Yesterday’s initial tests with the printer seems to have worked out fine. The rudimentary figures I tried to draw show up from the plates. Open questions about preparations for today were also answered by Maarten.

One of yesterday’s open questions was the amount of ampicillin that should be added both the liquid culture and the LB agar. It’s clear now – the final concentration should be 50 μg/ml, and thus I have to calculate the 250mg/ml concentration of ampicillin accordingly. Thus, the preparations I’ve done today included, for the liquid culture:

• 100 ml liquid LB
• 100 μl of our ampicillin solution
• and a swipe of bacteria

For the plates I made:

• 1l of LB – agar (0.75% agar)
• 2ml of our ampicillin solution
• 50ml of overnight liquid culture of the bacteria

Let’s hope that the way too low amount of ampicillin added yesterday still makes things work for now, as I’ve printed all 10 plates today, that were prepared yesterday. Another risk is that I might have mixed the bacteria with the LB agar while it was too hot. Unfortunately there’s no other way than to guess by hand if the LB agar is cold enough…

Today I tested printing with a wide variety of drop sizes and drop distances. We’ll see tomorrow which ones work out the best, if the plates are any good

A lot of preparation has already been kindly done by Maarten de Smit, who has arranged for the production of large quantities of LB agar, with 0.75% agar content (this is the agar content ratio of what is called the ‘top agar’). Trudie Bowers has provided me with all sorts of equipment and materials at the lab, like arabinose, liquid LB, pipettes, Erlenmayer flasks, liquid water bath, gas torch, large pipettes with rubber pumps, etc. Anne Kienhuis is organizing everything for me very kindly.

I will be working with 12×12cm plates to create my images.

The daily routine here will consist of maintaining a stable supply of plates full of bacteria in LB agar, so I can put them into the printer, and then experiment with the plates in the printer. To provide a stable supply, Maarten has grown some bacteria, taken from the cryogenic storage and spread out on a plate. Using this bacteria, overnight liquid cultures have to be made each day. Using these liquid cultures, the plates can be made themselves with bacteria, which also take overnight, and have to be put into an incubator.

Today I made a liquid culture with 100ml of liquid LB and some ampicillin. Here I’m in a bit of a trouble, as the ampicillin bottle says 250mg/ml, which seems an extremely high concentration, about 5000 times more concentrated than the 50μg/ml carbenicillin I used to use. Thus I put 1μl of it into 100ml of liquid LB, which should be 250μg of ampicillin. And of course a scrape of bacteria from the plate with the spread out bacteria.

I also made 10 plates, 12×12cm each with LB agar and liquid cultures. Used 1 liter of LB agar (0.75% agar), and put 50ml of liquid culture into it. I had the same dilemma here with the ampicillin, so these guys also got only 1μl, which should be 250μg again. I wonder if it works.

Afterwards, I started to fire up the printer and print on plates prepared by Maarten. At first of course there were some problems, but at the end it seemed to work out. Well, we don’t know of course – I got reasonably small drops of arabinose, but we’ll have to see tomorrow if they work out. Now I’m using a 16×16 pixel grid, which is old-school icon size.

• re-grow the bacteria on top of a plate, taken from the cryogenic storage
• create overnight liquid cultures from single colonies of the regrown bacteria
• create agar plates with the liquid culture in them
• perform arabinose experiments with it

Moreover, the UV-light bacteria killing experiment had to be re-done, as most probably the plastic cover of the petri dish itself filters out the harmful UV lights So I just did that, and have some plates exposed to UV lights without the petri dish cover, and now waiting in liquid LB to see if they grow out.

In more detail:

day 1: re-grow bacteria

Re-grow the bacteria on top of a plate, taken from the cryogenic storage. I did this on two plates today, one with just ampicillin in it, and the other also with arabinose, to see if it turns fluorescent at all. Easy reassurances create a flow experience

day 2: create liquid cultures

Take single colonies from the plates that grew overnight, and create liquid cultures from them. Also verify that the plate with the arabinose in it turned fluorescent.

day 3: create agar plates & perform arabinose experiments

Take the liquid culture that grew out and crate agar plates with the bacteria in them. As the plates are a few (at least 3) hours old, drop 5μl droplets of arabinose on them, and leave overnight to develop. Do the experiment in the following three settings:

• always have the plate in a 37°C incubator
• always leave the plate at room temperature
• always put the plate in the fridge

Moreover, have two kinds of plates for each experiment:

• regular LB-agar plates with bacteria in them
• ‘top agar’ plates

day 4: verify fluorescent pixels

Take the plates that had the arabinose dropped on them, and verify if any of the drops turned fluorescent, or the whole plate turned fluorescent.

Having exposed bacteria inside LB agar to 254nm UV light for 1, 10 and 100 minute durations – they still live. Whereas they should have died instead.

The other experiment revolved around plates with bacteria inside LB agar, left to grow at room temperature, first at room temperature and then put into the fridge 3 days later, or put in the fridge immediately. After 3 days, I dropped 5μl drops of arabinose on them so as to turn on the GFP-producing gene – but nothing happened.

So now I prepared some new plates, and also had one of the plates under 254nm UV light for an overnight time period. Let’s see if they work out…