Amazing how dramatic a single place can be in drylands in a season with some rainfall. Photos courtesy of Dr. Mike Westphal.
April 5th is the Graduate Symposium on Science Communication, with a poster presentation! I’m going to present a poster on the Baseline Cactus Survey completed in January, but I could use some advice. Let me know if you have any thoughts!
What are the cacti?
For the bird-cactus double mutualism project, we had planned on observing two study species: Cylindropuntia anthrocarpa (Buckhorn Cholla) and Opuntia basilaris var. basilaris (Beavertail Cactus). We also needed 3 class sizes (small, medium, and large) in which to bin the cacti. This would impact our sample size and equipment list. That being said, the best laid plans of mice and men (and grad students) often go awry. I’d only briefly visited our study site the summer before I’d officially started at York, so we knew we would need to revisit Sunset Cove to do some preliminary exploration before getting into the trenches and collecting end-game data. Getting to the site, it was immediately apparent that we would need to examine our plants more closely; there was nearly no beavertail in sight. So we altered the protocol, then added Cylindropuntia enchinocarpa (Silver Cholla) into the mix. The goal? Determine the location, size, size-variability, and health. We want a tall-ish species (so pollinators and frugivores would be interested) with plenty of variability in size, enough of them to manipulate conditions, and healthy enough so we can expect some flowers and fruit later on. And, for fun, we took a quick look at shrubs to see if they’re associated with cacti in any respect (I don’t go into that here, but the data is available on Github).
Where are the cacti?
Let’s make a quick map and take a look at the cacti individuals sampled. For C. anthrocarpa, we were easily able to sample at every 5 meters along 5 transects that were spaced 5 meters a part (n=105). C. enchinocarpa, however, was more sparsely distributed. So, after doing our first two transects 5 meters apart, we realized we needed to increase the distance between transects to 10 meters. We also weren’t able to get a cactus sample at every 5 meters, so we sampled 9 transects in total (n=98). The least common species, however, was Opuntia basilaris, which was so rare that transects were ineffective, so we instead unsystematically searched the entire site only to find a paltry number of individuals (n=26).
Based on the proposed protocol, we need 150 individuals of each study species to replicate each combination of variables 10 times. Ideally, the individuals manipulated between flowering and fruiting season will not be resampled in the the fruiting season, as our manipulation of the flowers in April may impact the number of fruits in August. This means that C. anthrocarpa is a solid study species option. C. enchinocarpa is certainly possible, but not as dominant as its cousin, and O. basilaris is out of the question.
How big are the cacti?
We’ve seen the distribution of cacti, but size of the cacti is what’s really important for this study. We need to know if the sizes are variable enough to split into 3 class sizes (small, medium, and large). We also need a general idea of their height to consider if pollinating and frugivorous birds will engage with the flower and fruits of the cacti at all. The three species did indeed have significantly different mean heights (Kruskall Wallis Test, p > 0.0001, df = 52, x^2 = 151.52), with means of 1.04, 0.55, and 0.17 for Cylindropuntia anthrocarpa, Cylindropuntia echinocarpa, and Opuntia basilaris, respectively.
How should we bin the cacti?
One important variable of our project is size classes within a species: small, medium, and large. Because height is what may influence pollination and frugivory, we will use the “z-axis” that we measured as the factor for size. Each size class must contain enough individuals for replication. We need to decide how to bin the size classes; either we can use natural breaks present in the data, or we can create equally-sized bins for the study species. Let’s examine each species’ size distribution, and make decisions about size class breaks on that.
None of the species have distributions with natural breaks (see density plots), and, especially for our two Cylindropuntia species, we can see that there are even distances between quartiles (see boxplots). For these reasons, I propose an equal-size binning method to determine size class.
Size-classes of cacti
But what exactly are the equal size classes for each species?
|Cylindropuntia anthrocarpa||<85cm||86cm – 152cm||>153cm|
|<45cm||46cm – 72cm||>73cm|
|Opuntia basilaris||<15cm||16cm – 22cm||>23cm|
We can see that Buckhorn Cholla (C. anthrocarpa) has the largest class sizes, followed by Silver Cholla, and then Beavertail. Having large classes may translate more clearly to birds, and therefore be a suitable metric to see if bird visitation is influenced by cactus size.
Health of cacti
Another important factor to consider when exploring potential study species is their overall health. After all, are these individuals even capable of flowering and fruiting? To measure health, we created a health index based on the Wind Wolves Bakersfield Cactus Report, which classifies each individual’s health on a discrete scale of 1-5 (1 being the least healthy, and 5 being the healthiest). We considered overall paddle/branch death, as well as scarification and rot.
We can see that the Cylindropuntia species are healthier than their Opuntia counterparts. The question is, will an unhealthy population still flower/fruit as much as a healthy population? Perhaps, but this is not the question of my project.
Who is America’s Next Cactus Superstar?
Considering its abundance, size, and health, Opuntia basilaris is not a realistic contender as a study species. It is likely to be overlooked by birds, not bloom/fruit due to poor health, and is in small supply. Therefore I must remove it from the running. Both of Cylindropuntias are healthy. Silver Cholla, however, is still less dominant than Buckhorn Cholla, is smaller overall, and doesn’t have the width of size classes that Buckhorn Cholla does. While these traits do not mean the Silver Cholla could not be a viable study species, I propose that focusing more on Buckhorn Cholla by deepening the methods of observation (i.e., joy sampling: stationary versus mobile count data, and increased hours of focal observation) will be more beneficial to answering my study questions than a comparative study between cacti species would.
This fall I have been processing the insects and pollen samples that I collected this spring from my fieldwork in the Mojave Desert. The insects were primarily caught using pantraps, and were transferred into 90% isopropyl alcohol for preservation. With the help of our lab’s two undergraduate practicum students, Shobika and Shima, we are gradually getting them nicely organized into collection boxes.
I pinned many, many bees and wasps when I worked on a pollinator census during my undergrad in West Hamilton. These are the steps I use for processing insect samples:
- Remove insects from alcohol.
- Give the bees a rinse in water to fluff out their body hairs (this step works variably well, we may need to give some of the larger specimens a spa day in the future)
- Gently dry with a paper towel, this causes the wings to uncurl. Wing venation is very important for identification.
- Under a dissecting microscope, pin from top to bottom through the upper right-hand side of the insect’s thorax into a stryofoam block. You want the insect to be completely horizontal.
- Gently uncurl the legs from the body and unfurl 1 antenna.
- Affix an insect identification label underneath the insect with the text readable from the left side of the insect. These labels should have date and location of collection, unique identifier and the name of the collector.
- Place into foam lined box.
- Very small insects get pointed rather than pinned. The right side of the thorax is glued to a triangle cut out of cardstock, and the triangle based is pinned instead.
I have also been mounting pollen samples whenever I can squeeze the time in. I collected stigmas from the field and have been storing them in ethanol-filled small tubes.
- Let slide warmer heat up
- Using a transfer pipette, remove the pollen-ethanol suspension and transfer drop by drop onto warm slide, letting the alcohol evaporate and ensuring it does not run over the edges.
- Place stigma onto slide.
- Rub the inside of the centrifuge tube that was storing the sample with a bit of fushcin jelly, place onto slide as well. Cut out 2 more small cubes of jelly, place over drop locations. Cover with slide cover and leave on warmer to melt jelly. Label slide.
For a different experiment that I have not yet processed, I will put the tubes into a centrifuge, spin down and pipette out the pellet to save time and labour. Quite a few tubes from the current experiment are extremely small and I am concerned about their ability to hold up under the force of a centrifuge. I need a less labour intensive process to make slides for my upcoming field season. I can think of two main options right now – use sturdy tubes that I can centrifuge, or collect into small tubes without adding ethanol, and mount each evening while at the research station. This will cut down the need to let the alcohol evaporate.