How to lose your eyesight

I’d love to be writing up my latest research for publication right now, especially since it's Academic Writing Month. But that project is currently in the waiting-for-various-things-out-of-my-control stage, so I can’t progress. Instead of biting my nails to the quick and sending inappropriately desperate emails across the globe, I have shifted focus to some lab-based tasks. What are these things? Come along and I’ll show you what fun I have! Both tasks I’m working on this week involve squinting at tiny things.

Task 1: Are the annual growth bands in my coral cores really annual?

Much of what I do involves collecting core samples from large coral heads. Much like trees, corals grow larger with time and form annual bands within their skeletons that can be visualized using x-rays or CT scans. I then measure the width and density of these bands to calculate the coral growth rate over the length of each core, and this tells me essentially how healthy the coral was over that time period.

(A) collecting a core from a nice big coral (B) the top of the core (C) what (B) it looks like once cut into a slab and (D) an x-ray showing annual density banding. If the top band in (D) is 2006, you can count years backwards as you move down the core, going back in time.

But of course this whole concept is predicated on the idea that the bands I identify are formed yearly. In some corals the banding is clear and lovely and life is happy. My most recent cores are not this type. They have painfully vague banding, and while I’d like to think that my experience means I can successfully identify the bands despite their lack of clarity, I’d like to be sure. 

This coral has nice banding. I like it.

This coral has rather shitty banding, and makes me want to poke my eyes out.

So, what to do? I first started by trying to count the number of something called “dissepiments” in the images. You can picture a coral as a tall apartment building, one that is constantly under construction; the coral adds a new floor to the top of the building once a month. Only the top floor is occupied by living coral tissue, hard at work on construction—once one level is complete, the coral seals this off and moves upstairs to start work on another. This is a decent illustration, because the coral skeleton actually looks a lot like this on magnification. The “floors” of the apartment building are equivalent to the dissepiments. All this is to say that one way to verify whether annual bands are annual is to count dissepiments—if there are about 12 of them for each of the bands identified, you are probably on the right track.

This could be easy if the corals behaved. (Nothdurft et al. 2005)

But really they look more like this and it hurts to find those little things the red arrows are pointing to.   (From Barnes and Lough 1992)

Another method is to measure the chemistry of the coral skeleton. While the coral is constantly building its skeleton, the composition of the skeleton changes ever so slightly with changes in the surrounding water—whether due to seasonal fluctuations in temperature, sediment in the water from river runoff, etc. I can use this to analyze a particular aspect of the skeletal chemistry that I know changes seasonally every millimeter down the core. This way I can put an independent time-scale on the core and then compare this with the time-scale I got by picking out my bands. In this case, I’m using the ratio of strontium to calcium, which changes due to water temperature (the skeleton is mostly made of calcium carbonate—CaCO₃—but other elements can substitute for Ca).

This is the idea. The black wiggly lines on the left show seasonal water temperature change (low in winter, high in summer), and conveniently the banding in the coral x-ray lines up with the wiggles! (From Bagnato et al. 2004)

Making these measurements is pretty straightforward but takes a lot of time:

(1) I cut the cores with a rock saw to produce a flat slab.

(2) I take the slabs to a medical facility and get them xrayed to reveal the particular convolutions of the coral growth direction in that sample.

(3) I further cut the coral slabs so that the maximum growth axis is exposed for sampling.

(4) Using an automated CNC milling machine and a lot of swearing, I grind a ledge into that exposed edge from which I’ll collect my samples.

(5) I clean out all of the powder from cutting the slab and milling the ledge that has accumulated in the coral’s pore spaces using an ultrasonic probe. This device blasts high-frequency waves through water such that tiny air bubbles form and explode, which helps clean the material, and destroy your hearing (I do wear earmuffs for this).

(6) The samples dry overnight and then I mount them on the CNC machine again and mill precise, tiny amounts of coral skeletal powder every 0.5 mm down the edge of my clean and beautiful skeletal ledge. Each of these bits of powder is caught on a square of waxed paper and then carefully transferred into a tiny plastic vial, labeled with the sample number. Too much coffee is not good for this step.

I get really excited when step 6 is over. Especially when I get to use pretty vials to spice up the lab-life.

(7) I acid-wash and dry a lot of larger plastic vials.

(8) I use a micro-balance (a very tiny and sensitive scale) to weigh out about 50 milligrams of coral powder from each of the 0.5 mm-increment samples into my clean vials. I attempt not to sneeze while doing this.

(9) I tire out my thumb using a pipette to add super-clean acid to each of the vials to dissolve the coral powder to the correct dilution.

(10) I gratefully hand the samples over to my colleague, who uses a machine called an Inductively-Coupled-Plasma-Atomic-Emission-Spectrometer to measure the Sr/Ca ratio in each of my dissolved samples.

Task 2: What’s up with the benthic foraminifera in my sand samples?

Foraminifera are single-celled marine organisms, and the “benthic” descriptor means they don’t live in the water column, but instead on the ocean bottom or on other substrates, like seagrass. Forams make complex shells, and in some areas these shells make of the majority of reef sands.

I’ve been collecting sand samples from my study sites and using a stain called Rose Bengal to dye all of the living foraminifera a lovely shade of pink. The dead shells remain white. This means that I can compare the living and the dead assemblages—the ratio of different types of foraminifera—to see if there has been a change over time (well, between “now” – alive, and “before” – dead).

Some of my pretty forams. This is from my recent paper with Sheila Walsh 

                                                                    

Benthic forams are sensitive to water quality—some types (A and B above) take over in dominance when the water is clear and low in nutrients, but this balance shifts (to critters like C-F above) if the water becomes nutrified (i.e. we dump sewage, fertilizers, etc. into it, or change the nutrient dynamics by removing the big tasty fish).

So how to quantify the assemblages? I scatter a scoop of my stained and dried sand sample onto a gridded tray, place it under my microscope, and then use a pin with a bit of surf wax stuck to the end to grab individual foram shells out of the sand and stick these onto little slides. Once I have enough, I count them. Voila! 

Unfortunately for my eyes, it takes a very long time to get enough—several hours per sample. And for science’s sake, I have a lot of samples…so…back to it!