Week 8: done and done

Hey everyone! So, it's been an eventful 8 weeks, but an extremely productive one.  Thanks for supporting me on this research expedition!  I will be leaving the island this coming Monday, to resume my day job as a curriculum designer for Iridescent.  This blog will turn into my personal blog and homepage, so I encourage you to stay tuned if you're interested in hearing more about science, education, online learning, and games.

To wrap up- first, if you donated at least $100 and would like a particular algae pressing sent to you, now's the time to say so!  I will be sending out the pressings this weekend, so please make your choice by Friday at midnight, otherwise I will select one at random for you.

A quick summary of the results of this summer:

I built a shear flume.  Yay!  It's super sweet and somewhat surprisingly works the way I intended. 

I got a little bit of data showing that the spores of one of the local species are as fast and as strong as the species in my thesis.  Thus the pattern seems general- red algal spores can produce an glue which can set, underwater, in a matter of minutes and is extremely strong once set.  We as humans have yet to manufacture a glue which acts nearly as good as this.

I also found that pH has a negative effect on algal spores- it seems to make them take longer to attach to surface, maybe making it more difficult for them to settle and become adults.

The research doesn't end here!  Becca Guenther, a PhD student at the University of British Columbia, has been following me around for the last few weeks, learning the ins and outs of the flume.  This research (in particular on ocean acidification) will continue as part of her PhD research.  I was really excited to hear her tell me this!  It means all the work I put into this summer will continue to develop into more and more interesting science, as the flume becomes a staple piece of equipment in the lab here at FHL.

I just wanted to end by saying thanks! to everyone who donated money and helped support my research.  I hope the blog was enjoyable, feel free to drop me a line or leave a comment if there's anything you want to hear more about or see.  Thanks again for supporting science!!

Kevin

Week 8: Graphs

Ok!  At this point, I've collected data on the attachment of almost 600 spores. That's what nonstop data collection for 2 weeks straight gets you. That's a lot of data points!  I finally got to plotting this data to share here.  This data is super-fresh, like 2 hours since I generated it.

The main species I've been using is Pterosiphonia bipinnata, (which I initially misidentified as a Polysiphonia). Here's what it looks like under a microscope:

 

It has these cute little tiers of cells that give it a banding pattern.  The spores are made on the inside of the tube of cells, kind of like a jelly-filled long-john donut.  You can see the packet of spores as lines of darkly colored dots.  These packets break into 4 individual spores once released.

Ok, and now to the main story of spore attachment.  I'm plotting the shear stress (or how strongly spores have attached) on the x-axis, with the time that the spores had been settling on the y-axis.  Since what I see is spores that are around at one shear and then gone at the next, I know that the attachment strength of the spores lies in between the forces of when I last saw it to when it was gone.  So I decided to plot the data as a bar, showing the range of where the spore's attachment strength must be, for how long they have settled.

This data is actually really cool!  It confirms the work of my thesis that spores attach really quickly and strongly.  How do I know this?  Well, notice that most of the points group on the left side of the graph, at really low attachment strengths where they are barely attached, or on the right side of the graph, where spores are really strongly attached.  I rarely catch spores at intermediate attachment strengths, meaning spores really just have one of two states: 1) unattached and waiting for a wave to knock them to a better location, and 2) fully attached, unable to be shaken by the strongest waves.  For Pterosiphonia bipinnata (as well as the species in my thesis), it may take several hours for them to "decide" to switch from state 1) to 2), but when they decide to do it, the switch happens faster than we can usually catch in our shear flume (on the order of minutes).

Oddly enough, Pterosiphonia bipinnata takes quite a while to make the decision to attach.  It is rarely attached in anything less than 12 hours, and even after 24 hours most but not all spores have moved to the right side of the graph (indicated by the darker color of the lines at 24 hours).  

At some point, I had accidentally tested a different species, and hadn't noticed till afterwards, when I had an outlier where spores were fully attached in 10 hours in just a few tests.  Going back to them, I found I actually had tested Polysiphonia hendryi. Here's the few data I collected for it:

And last, I ran a few pH tests.  So the big question: how will Pterosiphonia's spores fare in a future, acidic ocean?  The answer: not well.  In 24 hours of attachment, 73% of the spores had attached in the normal waters, but only 35% managed to attach in the acidic water.  It's interesting that as far as I could tell, when the spores did attach, they were just as strong, but seem to be taking longer to make that decision to attach.  I don't have enough trials yet to be fully confident in this story (I still got a few more days here!), but the trend so far is quite strong and I hope it'll hold up to more data.

Week 7: more data

Hey,

Quick update this week, as I'm busy plugging along for more data, now that everything seems to be working dandy (minus a power outage that happened this weekend, and set everything back a day).  Trying to analyze the data as I go, but haven't quit caught up yet.  Hoping to psot some results here by the end of the week.

Also, put a few more pressings up on the page, from the big stuff I collected from the FHL dock.  A few didn't come out so well, but there's still several good ones, if you are looking for the big and busty designer algae.  There'll be another 3-4 pressings added to the list by the end of the week, and then I'll ship them out the week after.  Claim your algae soon!

Week 6: DATA!!!

The last few days were a glorious ride. After struggling with many different species, I finally got on of the local fuzzy red species, Polysiphonia sp. (I'll key it out later), to release spores!  And then I saw the spores with a microscope, blasted them with my flume, and collected some super sweet images. The moment of first data is always a momentous occasion in any research project. At the moment I realized that the last five weeks of effort were going to finally work, I literally screamed "Yes, data!!!" at the top of my lungs.  Becca, who was helping me set this up at the time, looked over at me with an expression that was a mix of "Oh my god this guy is crazy," and "Yeah, I totally understand that feeling."

I found the only way to truly get really good images with the microscope was to eliminate any sort of air/water interface (discovered accidentally, as my microscope casing broken and filled with water).  So I needed to dunk the lens in the water.  I sealed the top of the lens so no water would get in the microscope barrel itself, then dunked the lens in water, and behold!: beautiful spores.  This was Moose's suggestion, and will probably ruin the lens at some point, but for now it's working splendidly.  But replacing a lens every so often isn't the worst operating expense.

Here's a with and without a barrier image, of the same set of spores:

And then, a sequence of image showing the number of spores remaining on the slide after hitting the spores with greater and greater force.  The first picture has 4 spores, which got knocked down to 3 and then 1.  Can you find them?

And last, the data graphic that resulted from this first set.  I had eight spores in view on this test, so I marked the information pertaining to the spores when they were de-attached.  So when the spore became deattached, how long had it been settling, and what force was needed to deattach it.  The points on the farthest right actually never became deattached. The darker points are actually two overlapping data points.

And as a final bit of news, I'll be posting some more algae pressings that are almost dry in the next few days. I got some really big ones in this set from the dock at FHL, after these I'll make one more set of smaller, more delicate (and in my opinion) more beautiful algae.

Week 6: Too stressed to spawn, and pressing highlight

Well, after recovering from a less-than-productive holiday week, I've returned to find my project still beset with problems.  There are only two problems now getting between me and data.

1) Seeing spores with my microscope.  This problem is mostly solved, I've now totally revamped my microscope setup, allowing for more maneuverability and clarity (a fancy way of saying I wrapped the microscope in a plastic bag and secured it with a rubber band). This hack job setup actually solved a whole bunch of problems- easier to light, more stable attachment, easier to maneuver, and better clarity. And for a brief moment, I actually saw spores! Three lonely spores, from one proud Calliarthron tuberculosum mother.

The setup is still a little unstable, and the vibrations in the carriage might actually cause blurry pictures when I use it to move the camera back and forth.  This problem is fixable though, either by 1) manually moving the webcam to set positions, 2) making a new carriage or 3) or if all else fails, counting the spores by eye, without the webcam. I will figure out which solution to use when I have spores to look at, which leads to the next problem.

2) No spores.  I've now confirmed that the algae I've been working with have just not been releasing spores, or releasing in such low numbers as to be useless.

I've been mostly working on Corallina vancouveriensis, the same species I worked on in Monterey and had good success with.  After several sets of no spore release, I finally took them under a microscope and dissected their reproductive structures yesterday.  And, well, they didn't look so hot.  Compared to Monterey, they had lower numbers of of reproductive structures, lower numbers of spores in their structures, and the ones that did have spores had pretty crappy, deformed spores.  So, no wonder I wasn't getting good release.

I talked things over with Becca, and we formulated several strategies:
1) Collect Corallina in a new location on the island, where it might be healthier.
2) Get some divers to collect Calliarthron tuberculosum, my other study species. This species also doesn't look too sexy in the intertidal, but Becca said she's found healthier and more reproductive fronds from subtidal locations.
3) Put intertidal Calliarthron in a dark growth chamber, where a colleague of hers said she got that species to form reproductive structures.
4) Switch to an entirely different coralline or noncoralline species, chosen based on local health and reproductive state.

We're actually going to pursue all these strategies simultaneously for the rest of the week, hoping at least one of them pans out and solves this problem.

Pressing Highlight: Microcladia borealis

This highlight features a relative to one of our previous favorites, Microcladia coulteri.  The second and only other representative from this genus in the area, Microcladia borealis looks a fair bit different from its cousin.  It's less flattened and has more tube-like branches that often become entangled with one another, causing it to look like a hairy mess in the field.  But pull it apart for a pressing, and it's simply gorgeous.

The species has what's called a unilateral branching pattern.  That simply means when branches form off a trunk, they always form on the same side.  Those branches can form sub-branches, but the sub-branches will again pick a side and only form on that side.  You can see this unique pattern in the pressing.  This gives a unique design and shape to the algae. I always thought this algae would make a great suit in a deck of cards, like a club or spade, or frond (hehe).

This algae is generally pretty small, with branches extending usually no more than a few inches from the main branch.  But because of it's delicate size, it takes extreme care and effort to turn it into a pressing.  I spent more time arranging pressings for this species than I do for any other.

Week 5: if only I could see...

Vision issues.  Everything is basically complete, except... I haven't seen spores yet.  And this continues to be a problem.

A few test runs this week got algae to release spores in my tanks, but that I simply wasn't seeing them.  After scourging the microscope and webcam optics all day yesterday, I finally found the culprit: bubbles, and maybe low light levels.  Because the my tanks are constantly bubbled with CO2 enriched air to keep the acidity constant, there are just too many air bubbles floating around.  In almost not time flat, they cover the glass plate under my microscope and just make it impossible to clearly see anything

A picture of the setup, in total.

I think this is a solvable problem, I'm just figuring out how.  I improvised a plastic bag to contain the microscope today, and it seemed to work pretty well. Further tests with spore release will tell. It's really annoying to be so close to my first really data, and to have bubbles get in the way.

In any case, went on another beautiful collecting trip.  I got some great pictures of the three species I'll be working on this summer: the two pink corallines from my PhD, and a third fleshy (not calcified) red algae that I've had good success in the past getting spore release from.

Corallina vancouveriensis, the one that look like rose petals

Calliarthron tuberculosum, spiny and spickly.

Polysiphonia sp.  Really common, mat-like algae.  Pull it apart and its a bunch of fine, interwoven branches.

In any case, happy 4th of July everyone!

Week 4: Flume calibrate and Pressing highlight: Porphyra sp.

Week 4 has ended on a highly positive note.  I spent most of Thursday and Friday afternoon working on calibrating the flume (Friday morning was spent on a lab kayaking trip!).

I built a device called an anemometer, which used glass tubes to measure the amount of pressure caused by fluid motion. I used it to measure the pressure at several points along my base plate.  By knowing how much pressure drops along the base plate, I can directly calculate the amount of shear force that is causing that pressure drop.  And thus I know how much I will be blasting the spores in my tests.

I was so wrapped up in the testing, that I didn't take any pictures of the setup!  But I do have the first data graph to show you.

This shows how the shear force that I'm generating corresponds to the height of the pipe I use in the flume.  The trend is pretty linear, as hoped.  If I fill up a pipe twice as high with water, then twice as much pressure is driving water through my flume, and I get twice as high shear forces in my flume.

And I was able to hit 30 Pa, the magic shear stress number. For whatever reason, most studies haven't been able to go much higher than 30 Pa, so as long as I can get to that force, my flume is working as strong as anyone elses.

I did learn a few things from this testing, such as how the position of my clamps can effect the forces I measure by allowing water to leak out the sides of the flume.  To compensate for this effect, I had to adjust from a four clamp system to a ten clamp system, meaning I had to go buy a bunch of clamps from the hardware store.

Pressing Highlight: Porphyra sp.

This week's algae is probably the most famous of seaweeds. It comes from the genus Porphyra, and is also know as nori, the stuff you wrap around your sushi. It's a red algae, paper thin and quite delicate.  To make nori, Porphyra is ground up, layed flat, and baked into a textured sheet.  You can eat Porphyra (as well as many other algae) raw, but I'll confess it's much improved by the nori processing.

You'll notice that I've left the description at sp. (meaning unidentified species) rather than giving a species epithet. I know there a few Porphyra species in Washington, but I'm not that great at telling them apart. From talking with Porphyra experts at Phycology meetings, I've learned that even the experts are still creating and redefining new species constantly.  So even if I were to use a guide to find the "name" of this species, there'd be a decent chance that name would be wrong anyways.  As a results, I don't bother and just leave most species in this genus as Porphyra sp.

The species I've pressed look a little beat up, almost as if their degrading. But the degrading is perfectly normal for this genus, the red degraded regions are the reproductive parts of the plant.  Porphyra gets reproductive on it edges, converting it's tissue entirely to sperm and eggs.  Once the reproductive tissue is primed and ready, it simple deattaches from the parent plant, to float in the water and find a mate! The reddish color indicates that this is a female, males have white edges.  Some of the red, gooey portions around the plant were egg packets about to be released, and got caught around the edges of the plant in the pressing process. So actually, this individual is quite sexy at the moment, and we quite literally caught it in the act in this pressing!

Week 4: Testing and tweaking

Halfway through week 4, and moving along.  I solved the last two problems I was worried might trip me up on Monday, making sure the cameras could actually focus on the surface of the glass and weren't too far away, and getting enough light to a microscope immersed in a container of seawater so that I could actually see things underwater.

It's always the little details like this that you don't really think about until the last minute, when you are hooking the webcam into the camera and thinking "So this is going in a tank of water- where's the light coming from?"  And then you hope you're creative enough to find an answer to a problem you didn't realize existed until moments before, but might wreck your whole experiments. So far, I've been lucky/creative enough to not have anything permanently trip me up yet.

I tried a trial test run on Tuesday, where I let an algae release spores and then tried to take pictures of them with the webcam.  It seemed to work, but I never found the spores with the microscope.  They either never were released, got blown off when I assembled the flume, or (most likely) were just too dang small and sparse to find on my giant sheet of glass. I think with a little more care, I should be able to find them, hopeful this won't be the devilish detail that wrecks the whole project.

In any case, here's a picture of my more or less finalized setup:

I made markings to tell where I was on the glass:

And was able to see the marking through the webcam, onto the computer:

Also, this little guy is my favorite example so far of creative troubleshooting.  I was having difficulty finding a way to secure the motor to the metal framing of my setup.  After trying a couple very clean and official techniques, I started getting sloppy and just using zipties to hold it down so I could temporarily use it (and it just so happened the nearest zipties were bright pink).  Finding that worked, I started adding more zipties. And more. And then I laughed maniacally to myself as 8 zipties later I had strapped the motor in every direction I could think of, and by golly it wasn't a pretty solution, but it worked!

Week 3: Pressing highlight- Microcladia coulteri

Hey!

Good news, Becca got back on Thursday, we were able to hit the low tide on Friday, and I got some algae pressings done this weekend!  No pictures from the field as it was a pretty crappy, rainy day.  But I wanted to introduce the algae I'll be pressing with short algal vignettes. Hopefully these short stories will give the algae pressings a little more character, and help you appreciate algae the way I do.

There's lots of seaweeds with lots of shapes, sizes, colors, etc., and I'd like as much as possible to be sure everyone who is receiving a pressing as a reward gets one they like. So as the algae pressings dry (which should take about a week), I'll post pictures of the pressings on a separate webpage, where people can claim them on a first-come-first-serve basis.

Week 3 Algal Vignette: Microcladia coulteri

Not all seaweeds are created equal.  Finding the right seaweed for a good pressing can be tricky, some algae that look beautiful in the water just become dull, pasty scraps when pressed. But not Microcladia coulteri: it looks good in the water and on the press.  It has a vibrant red color that dries well, a flattened, two dimensional growth that presses well, and a rugged cell tissue that can be easily spread out without breakage.  It's sort of a classic to me, like Old Spice or Chuck Norris. When I think about pressing algae in the Pacific Northwest, it's the first thing that comes to mind.  So I suppose it is appropriate to have found it on my first trip and to use it in my first pressing.  Microcladia coulteri isn't super abundant on the San Juan Islands, but if you know where to look it is common enough to be easy to find.  So in video game terminology, you might can it an "uncommon drop."

Week 3: tidying up

Most of the way through week 3 now.  Everything is going smoothly, pretty much on schedule.  I plan to start real experiments come this Monday.

I spent Monday and Tuesday working on coding for the Arduino/webcam setup, and amazingly figured it out.  Check it out, here's the solution I found: I program the arduino to move the motor a set distance, then stop.  I then use the Arduino's built in serial port (genius feature!) to send a signal back to the computer saying "Hey, I'm ready, take a picture!"  Meanwhile, I have a Processing code (new language to me, it's pretty sweet) running on the computer waiting for the Arduino's signal.  Once it gets the all go, it says "Ok boss, picture time," and tells the webcam to take a picture. Slather, rinse, and repeat, for a series of spatially organized pictures of baby spores triggered by the push of one button. Pretty sweet, huh?

Since then, I've been mostly finishing up odds and ends, connecting pieces and cleaning up joints.  Here's some pictures of the flume in a more or less finalized state.

Unfortunately, no field trip this week.  Becca, the grad student who will be working with me on this flume, is currently out of town and delayed in getting back.  She's also my guide to the local algal hotspots, so without her, a field trip is a bit pointless.  Oh well, more on the next low tide.

Week 2: The robot moves!

The last two days were a huge success! I managed to get a fully functioning carriage move back and forth, powered from a stepper motor using a code from an Arduino!  I'd like to say it was my ingenuity that solved my carriage problems, but most of the credit really goes to the lubrication properties of WD40.

This shows the carriage assembly in the coolers that I'll be using for the experiments.

 

 And, a video of the action:

Unfortunately this drama with the carriage has put me behind schedule a bit.  The flume is nearly assembled, next week will involve assembling the flume, figuring out how to take pictures with a webcam using an arduino, and final programming for the arduino code (not the most exciting or photogenic of activities).  I don't think I'll actually get to calibrating and testing the setup until week 4. 

On the plus side, I've now figured out how to make and bought all of the necessary components for all of the different aspects of this project.  Just in time too, I've nearly reached the end of the budget!

In other news, there's great low tides next week!  I plan on going on collecting field trip, and finding some nice algae to use for the first few algae pressings.

Week 2: the flume arises

And week 2 it is!  A lot happening here, the flume is starting to become something recognizable.  Here's a few shots of the pieces I've constructed so far, not much on their own, but they'll look impressive soon.

The base plate, the clamps are holding the glass plate to the plastic sheet until the glue sets.  The spores will settle on the glass.

Moose had some concerns my original design for the connector box being able to supprt the weight of the water and pipes.  So he had the great idea to turn my "box" into a "elbow" (or a 90 degree connector) which solves the problem of the weight support but still leaves me with a circular cross-section of water that need to be rectangular.  To solve that problem, I cut a rectangle out of the pipe cap, and am going to fit a few plastic pieces to form box that reduces the flow to the shape I need for the shear flume.

 

This is the piece that fits over the base place, and will connect to the connector box. I'm basically just gluing two plastic bars to a sheet of glass.

The biggest holdup so far is the carriage assembly, the thing that the motor moves back and forth.  This has proved both more difficult and more expensive than expected. Right now I plan to use the motor to spin a threaded pipe, with will force a block to move back and forth. But I bought a $50 carriage on McMaster (the key piece of the assembly) that didn't really move smoothly at all, meaning the threaded rod with probably get stuck rather than slip through.  I'm going to make some adjustments tomorrow and try to make this carriage work, but I might need to find another way.

The carriage assembly is also way more costly than I expected.  It seems like each assembly will run over $100, way more than I anticipated (I guessed $20), and even at that price, it's not all that great.  But then I realized that I am completely automating my data collection process, and that this is a pretty awesome and creative way to do it, and when it works it will be so worth it. I had to manually count spores under a microscope as fast as I could in my PhD, and that to avoid that again is worth a little time, energy, and money.  

In any case, it's definitely a good thing that I got more than I asked for initially.  Thanks everyone!  Your generosity has covered my cost underestimate.

Week 1: motor controls

Wow, eventful first week!  I spent more time than I would have liked getting settled, ordering final parts, and meeting various people, but I still managed to make some progress.

Arduino is ridiculously easy to use!  I sat down with the parts and software on Thursday morning, and by the end of the day, I was able to connect the arduino to the motor and use a code to make the motor spin!  Yay!

Sure, this isn't on the forefront of technology or anything, but it's still pretty damn amazing to me that I can write some random, arbitrary characters on a computer, push a button, and get a motor to do something.

It gets better though, because Moose let me borrow his Arduino LCD shield (a chip that connects to and sits on top of the arduino, giving you a simple display and several buttons).  But the end of Friday, I had programmed a neat little menu into the LCD display that let me control the motor completely through the Arduino.

And that was about it for real progress this week, though honestly, I didn't expect to get a nearly finished code in arduino so quickly, so this is actually more than I expected to get done.

I'm making a page with my current planned schedule, in case you want to follow along and know what's ahead.  Finally, I'll end with a sprinkle of few pictures from the ferry ride here, and the field trip on Wednesday.

Damn, I wouldn't mind living in that house.

Friday Harbor Laboratories, as seen from the ferry as we pull into the port.

Seaweeds! If you're not looking forward to many glamour shots of seaweeds, you're probably on the wrong blog.

A giant Pisaster seastar! And bigger than my hand!  Special exception here due to it's remarkable size, don't expect many pictures of disgusting animals.

Week 1

Ok! All packed and ready to go to Friday Harbor tomorrow morning, via bus, plane, bus, and ferry.  I'm posting this in fear of being one of those annoying bloggers who writes about every little thing that happens ("Now in the Terminal", "Now boarding the plane", "Now exiting the plane!").  Trust me, you'll get none of that.  I am pretty new to this blogging thing though, so if I start doing it wrong, feel free to let me know.

I'll aim for a twice-a-week posting schedule to keep everyone updated, without being overtly annoying. This week, I'm ready to hit the ground running! I'm flying in during a low tide week, so appropriately have some tentative field trips planned for this Wednesday and Thursday with Dr. Patrick Martone, a professor from University of British Columbia and coralline algae expert.  Also, he used to be my labmate and is a good part of the reason I study corallines now. He's one of the two professors I'll be working with this summer.

Diagrams!

Special treat today, for all of you eager to see what the hell this contraption looks like.  I sketched out a simple hand-drawn diagram.

As you can see, I'm no artist, but hopefully you get the idea.  There are four pieces, a base on which the spores settle, and then three attached pieces which when added to the base create a shear flume.  Water builds up in the gravity flume, creating pressure.  The water then transfers through the holding box, whose purpose is to connect the vertical 2" pipe to a thin narrow shear channel.  The cover fits over the base, creating a thin narrow opening through which water flows to create shear.


I've now ordered the raw materials to construct each of these components.  Yay! Turns out custom-cut glass isn't so hard to find on the internets, even found a place in Washington to supply the glass pieces, so it shouldn't take them too long to ship to Friday Harbor.

Arduinos!

Well, I jumped on the Arduino bandwagon.  Definitely coming late to this one, but it seems a hot topic right now (meaning, I have 3 friends that are really excited about Arduinos right now, one being Moose, the post-doc at Friday Harbor that I'll be working with.  Yes, he goes by Moose and you'll hear more about him).

For those still new to the bandwagon, Arduinos are the new wave of opensource, multipurpose microprocessors.  Yay!  What does that mean?

They are basically mini computers, about the size of a well-made S'mores.  They got a couple inputs and outputs, and as I've learned are great at controlling simple stuff, like stepper motors and your roomate's alarm clock on April first.

The best part is that they can be controlled by their own, open-source programming language.  So you can give them a set of commands, and they run like a super simple computer.  Open-source is key here as it aides laziness- you don't need to program it from scratch, but can use any one of the many programs other people have kindly made available all around the internets.

Only two pieces left to buy now, a moving platform to hold the microscope, and the glass plates for spores to settle.  Unfortunately I found out that McMaster-Carr (the number one online source for raw materials) does not sell glass sheets in 2 foot lengths, so need to find another option.

I swear one of these days I will upload schematic diagrams so you can understand how this all fits together.  Not now though.

First purchases

Alright, the project has started!  Put in my first orders for the microscopes and raw materials from the flume.   You can see them updated on the expenses page.

Already creating problems, as I'm waiting to hear back from McMaster on the price of custom-cut glass sheets.  I'd like to have a glass bottom for the spores to settle on, but if McMaster charges me through the roof, I'll have to settle for some scratch-resistant acrylic or something lame like that.  Fingers crossed!

Welcome backers!

If you got here, you must have backed my project, Mystery of the Tiny Algal Spore on Petridish.  Thanks so much for the support!  Through my project, I will post updates to the blog, so to follow along, I'd suggest subscribing to or bookmarking this blog.  For the first couple posts, I'll update on Petridish when I post here, but eventually I'll just post here.

A quick preview of the schedule: I will be spending the next month or so buying supplies and building prototypes for this summer.  Then on June 5th, I will fly to Seattle, where I'll catch a bus to a ferry and head to the beautiful San Juan Island for a month.  There, I will work in Dr. Emily Carrington's lab at Friday Harbor Laboratory to do the project I described on Petridish.  Finally, I will leave the island on July 31st and fly back to California, with hopefully a completed project