A workflow for pollen identification.


The reproductive ecology of cactus is not well-studied. A small, side project of mine is to determine the pollinator guild of buckhorn cholla at Sunset Cove, Mojave Desert, and with which plant species, if any, it shares pollinators. The genera Opuntia and Cylindropuntia are known to be insect-pollinated, but I am curious which of the more than 659 species of bees in the Mojave Desert desert are pollinators.

As visitation does not necessarily lead to pollination, I removed the pollen loads from 22 bee visitors I caught during insitu observation periods. I also removed stigma from the cholla to quantify heterospecific pollen deposition i.e. evidence of pollinator sharing. Pollen ID is not easy task and so I have developed a workflow to make it more streamlined.


Prep a reference collection:

  1. Create a reference collection by removing pollen from the anthers of several flowers of every species blooming in the area. Store in ethanol.
  2. Mount and stain the pollen with fushcin jelly.
  3. Image each species of pollen grain at 3 magnifications. Measure the length and width of about 10 grains per species. I calibrated Lumenera’s Infinity Analyze software using a stage micrometer to make this really quick.
  4. Make a reference document to consult. I use a word doc where every page is a species. Add in the photos at several magnifications, the mean size and any notes.
Sample reference page for Echinocereus engelmanni (Hedgehog cactus)


To go through the stigma or bee pollen load samples, I use my Canon EOS 60D dslr with a 60mm macro lens pointed confocally into a light microsite at 100x. I used the remote shooting utility from Canon to control the camera with my computer and display the view onto a second monitor.

Home example of confocal setup
  1. I designate each coverslip on the slide as a zone and do 8 transects through each, counting the grains. Each line in my spreadsheet is a transect, each column is a species. I use 5 columns for buckhorn so I never have to count very high.
  2. I don’t count damaged grains, or grains in air bubbles.
  3. Each slide gets its own folder. I take photos of each heterospecific grain with the file name as the zone + transect + species, which is simple using the photo utility. Knowing where the grain is on the slide and what its surroundings are will be helpful if you need to find it again.
  4. The species can be tentative for now so don’t get too bogged down.
  5. Take photos of unknowns when first encountered and assign them morphospecies ID. I put these in a separate folder as a reference.
  6. Some species are easy to ID. Quite a few are not. The more grains you see the easier it is to spot the differences.
  7. To help ID, we can take a page from entomologists. Sort the photos by their tentative IDs, putting each species in a folder so they are visible all at once (do a bulk rename to append the folder name first). It is difficult to compare grains unless they are side by side, which isn’t realistic with one microscope.
  8. Sort until each folder contains identical grains, then assign them a species from the reference collection. Or assign them to a species group for species that are virtually identical (likely Asteraceae!). Assign any remaining to morphospecies. Update the datasheet with the corrections.  
Buckhorn cholla (larger) and silver cholla (smaller). Thankfully the most abundant grains are simple to differentiate.

Field sample processing

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.
  • Repeat!
  • 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.

Process:

  • 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.

mini-reviews

Mini-reviews are shorter and more focused than traditional literature reviews. Their specific format varies between journals, however they all have a few things in common: They are topical, concise and specialized, rather than being exhaustive. They quickly bring the reader up to speed on current research in a field, particularly when there has been a major change in thinking. This is in contrast to major reviews, which provide a comprehensive overview of a subfield.

Mini-reviews often synthesize recent research, offering insight and new direction in an important emerging research area. They ideally propose new ideas and hypotheses that arise from the synthesis. Challenging current views in ecology and embracing a bit of controversy is welcome. Despite being called minor, these reviews may garner higher readership and impact than major reviews, due to their conciseness, readability and relevance. I think they are particularly suited to interdisciplinary synthesis, as they do not require writing an exhaustive background from each field, making it easier to communicate the interesting or important aspects of the crossover to a wider audience.

While only a handful of ecology journal explicitly provide guidelines for a mini-review, but quite a few impose a shorter word limit (< 3000 – 5000) and limit references to around 40, essentially requiring a mini-review. Other keywords I have noted are ‘topical’, ‘specialized’, ‘research reviews’, ‘briefings’ and ‘question-based’.

They following ecology-related journals either publish mini-reviews by name, have previously published mini-reviews or their submission guidelines strongly suggest that they welcome the format:

  • Journal of Ecology
  • Methods in Evolution and Ecology
  • New Phytologist
  • Annals of Botany (Botanical Briefings)
  • Frontiers in Ecology and the Environment
  • Global Ecology and Biogeography
  • Conservation Letters
  • PLoS Computational Biology
  • Insect Conservation and Diversity
  • Basic and Applied Ecology
  • Functional Ecology (question-based)
  • Ecosystems (invited only)

Mojave Desert Site Selection

I recently had the chance to spend a few days exploring in the Mojave National Preserve in California to select new study sites for my study of plant-plant-pollinator positive interactions. Sunset Cove, located in the UCNRS Sweeney Granite Mountains Desert Research Centre, is host to an incredible diversity of shrubs and cacti. The two foundational species I will be studying are creosote bush (Larrea tridentata) and buckhorn cholla (Cylindropuntia acanthocarpa), which are codominant in this site.

Annuals were germinating all over the site, however they are still incredibly tiny. Both creosote bush and buckthorn were showing facilitation, with higher abundances of germinants under the shrubs than in open areas. Finally, one cool observation was that 100% of the buckhorns surveyed were growing in close association with another codominant, but only 70% of creosote bush were growing with a codominant, suggesting cacti have some interesting interactions in this area. Finally, this area has been experiencing winter rainfall leading to optimism about flowering this spring.

 

 

Potential study species

The reproductive biology of Cactaceae is not well known – only approximately 2% of the 2000 or so species have been studied (Mandujano et al, 2010). Consequently, how they interact with neighbouring plants of different species for pollinators or what this means in a community context are both virtually unknown. In one of the few published experiments that explicitly tested these interactions, researchers focused on the highly invasive prickly-pear Opuntia stricta in coastal shrublands in Catalonia (Bartomeus 2008). Cacti in the Opuntia genus are primarily bee-pollinated; they have large, colourful bowl-shaped flowers and many species are rich in pollen and nectar (Mandujano et al, 2010), suggesting they are very attractive to pollinators. Plants that exhibit these characteristics can interact with other plants in two notable ways for pollinators – they may act as a magnet plant, increasing local abundances of shared pollinators and thus facilitating the pollination of their neighbours, or conversely, they may steal pollinators and reduce the fitness of their neighbours.

To determine the effects of the invasion on the native plant community, the researchers created plant-pollinator interaction networks for both invaded and uninvaded sites. They found that O. stricta acted as a super-generalist in its new range. It was visited by 31% of the insect taxa in the invaded sites and was outcompeting native plants for pollination services. Within the same study, they found that Carpobrotus, an invasive succulent, had the opposite interaction with the surrounding plant community; it facilitated the pollination of the native plants in the system. This highlights the species-specific and context-dependent aspects of these interactions. There are a few species of Opuntia common in the Mojave Desert, and I hope to discover if and how they are interacting with other plants, particularly shrubs and their annual understory.

Attribution Stan Shebs [GFDL (http://www.gnu.org/copyleft/fdl.html), CC BY-SA 3.0 (http://creativecommons.org/licenses/by-sa/3.0) or CC BY-SA 2.5 (http://creativecommons.org/licenses/by-sa/2.5)], via Wikimedia Commons

Bartomeus, I., Vilà, M., & Santamaría, L. (2008). Contrasting effects of invasive plants in plant–pollinator networks. Oecologia, 155(4), 761-770.

del Carmen Mandujano, M., Carrillo-Angeles, I., Martínez-Peralta, C., & Golubov, J. (2010). Reproductive biology of Cactaceae. In Desert plants (pp. 197-230). Springer Berlin Heidelberg.

Progress Report – Fall 2016

Several weeks ago I completed my first progress report for my MSc program. This involved a giving a short presentation (slides above) followed by a question/discussion period. My thesis focuses on pollination facilitation – non-competitive pollinator sharing between plant species that improves the reproductive success of at least one of the participants. I will be investigating these interactions in the Mojave Desert, a biodiversity hotspot supporting 659 species of bees and 1680 annual plants.

Why spatial? The study of ecology is normally separated into hierarchies, however, we know that these different levels are integrated and interact despite studying them in isolation. All interactions take place in space, and so explicitly including spatial dimensions to a study can be a way of connecting these levels, leading to a deeper understanding of the observed interactions.

It can be a little intimidating to stand in front of your committee and tell them your ideas, but they are there to support you. I received some great feedback which I am using retool my experimental design in preparation of the upcoming field season. Advice: Be careful about your clipart choices! I used a picture of queen honeybee (they don’t pollinate!!) in an interaction diagram explaining pollination facilitation. This isn’t as bad as the infamous biology textbook “Bees of the World” showing a pollinating fly on the cover, but it was noticed right away.