Thus- it is critical that we all take steps to prevent more damage to our oceans and at the same time help the oceans recover. The first step is reducing our waste, which I will cover in a future blog, but another step that we can all do is to cleanup our parks, streets and beaches to prevent more plastic waste from entering our oceans. We can even do this during Covid19- while socially distanced outdoors and wearing our masks and gloves!
So this ‘Zoomester’ I decided to organize a Plastic Cleanup- with an in-person beach cleanup event at Playa Del Rey Beach in Los Angeles and a remote option for those individuals that were living afar. This took place last Saturday (10/17/20) and all in all it was a tremendous success with about 15 people that participated! This included faculty, staff, graduate students and undergraduates, including four undergraduates from my Environmental Studies class that I had never met in person prior to this day (I get so excited to meet people in real life now.. ha). In the below photo are two students (and roommates) from USC posing behind some of the trash that they and several others collected. Unfortunately I forgot to ask everyone to stack their trash, so I wasn’t able to document all of it.
The fabulously talented photographer: Maurice Roper (USC) also came and documented the whole event! Below I have included a gallery showcasing some of the photos he took!
In addition to Maurice Roper documenting the event, I was very fortunate to have the wonderful support of USC’s Environmental Studies Program and the Wrigley Institute. I want to give a special shoutout to Dr. Jill Sohm (Director of the Environmental Studies Program) and Dr. Ann Close (Wrigley Institute, Associate Director) for their help. Lastly, I was able to use hands-free online waiver forms with the help of Kate Tucker at Resmark Systems with “WaiverSign” (I highly recommend them for your hands-free events!).
I truly felt like this event was impactful. Aside from all of the trash that we cleaned off the beach (the majority of which was plastic), there were many people that watched us and thanked us, and even some that joined us! So I have hope that this event spread awareness as to the little things that WE CAN ALL DO to help our oceans and environment!
Last Thursday on a typical sunny Californian day, I met up w/ researchers, Kirsten Sheehy (UCSB RIVRLAB), Bill Neill (Riparian Repairs) and Noa Rishe (California Department of Parks and Recreation) to romp around Topanga State Park in search of the invasive Cape ivy, Delairea odorata. As soon as we stepped foot onto the Los Leones trailhead we could see the vast entwined vines of this relentless invasive ivy climbing over and smothering all of the native species in its path, hence its nickname- the ‘California kudzu’.
Me, standing in a thick blanket of Cape ivy near Los Leones trail in Topanga State Park
Kirsten Sheehy, Research technician at the UCSB RIVRLAB inspecting the best place for the first release of the biocontrol agents- the gall-forming flies
Native to South Africa, Cape ivy was originally introduced into the USA in the 1850s for ornamental purposes due to its pretty green color with lush yellow flowers. However, looks can be deceiving as in some areas this invasive weed has reduced native plant species richness by 36%, and decreased native seedling abundance by 88% (see Alvarez and Cushman 2002). In addition to its detrimental impacts on native plants, this invasive weed also produces many chemical defense compounds (eg. pyrrolizidine alkaloids and xanthones), which make it toxic and unsuitable for foraging by resident mammals; and potentially detrimental to fish survival if substantial amounts of these chemical compounds end up in waterways. Aside from its toxins, this weed can interfere with nesting sites by many riparian-dependent birds. This invasive weed is also quite the ecosystem engineer due to its shallow root system contributing to serious soil erosion problems on hillsides; as well as potentially forming a serious fire hazard due to its dried out foliage hanging over native trees during the dry season.
Thus, there is a dire need to control the spread and growth of this menacing invasive vine. Invasive weeds can be controlled in several manners including herbicide chemicals, mechanical removal (via hand-labor or machines), and biological control. In classical biological control, a pest or weed’s natural enemies (for example, the insect herbivores of Cape ivy) are collected from its geographic place of origin, tested for target specificity and efficacy, and then released into the invaded region.
Successful biocontrol agents can reduce pest populations below threshold levels that cause problems for humans and native species. Once established, biocontrol agents can provide a sustainable, long-lasting management option as biocontrol agents are self-reproducing and self-distributing. Biocontrol agents will not eradicate every target pest or weed individual but this is actually a positive feature as it prevents population crashes of the biological agent and promotes the long term control of the weed. In sensitive or protected regions, biological control and hand-removal of invasive weeds are often the preferred method of control in order to reduce any negative impacts to the surrounding native ecological community.
Hence, last Thursday was quite a monumental day as it marked the first release of the biological control agent for the control of the invasive cape ivy in the greater Los Angeles Region. The biological control agent in this case is the gall-forming fly, Parafreutreta regalis Munro (Diptera: Tephritidae), that has already been approved for release after undergoing intensive testing through the USDA-ARS to ensure that it only targets the invasive cape ivy, in order to prevent any non-target effects on local plants. Similar to its host plant, this gall-forming fly is native to the Cape Province of South Africa and is known to stunt the growth of Cape ivy in both the laboratory and in the field. Thus, it is expected that this biocontrol agent will reduce cape ivy’s ability to spread and climb, both which would reduce the smothering impacts of this invasive weed on native vegetation.
In two weeks from now Kirsten Sheehy and the UCSB RIVRLAB will come back to make sure that the galls are forming on the ivy before removing the cage. Once the galls have formed, these flies are pretty much on their own, continuing the cycle of injecting their eggs into new unsuspecting cape ivy hosts, and forming new galls that promote generation after generation of weed-controlling superheroes. Of course Kirsten will continue to make periodic new releases of adult flies in the SoCal region to increase the genetic variation of this fly to ensure the success of these new populations in the Los Angeles Region. The goals are to establish these super-hero flies in at least one site per coastal county in California to serve as ‘nursery’ sites for future regional releases. San Diego Co. is next up on this lucky-list of biocontrol study sites.
In addition to this beneficial fly, further biological control research on a stem- boring moth, Digitivalva delaireae Gaedike & Krüger (Lepidoptera: Acrolepiidae), is underway via the USDA-ARS (see Mehelis et al. 2015) and is likely to be approved for release in the near future. This stem-boring moth is actually expected to have an even greater impact on controlling Cape ivy, especially if it is combined with the impacts of the gall-forming fly. Once approved, we hope to add this additional superhero biocontrol agent in the SoCal Region in order to reduce the ecological crimes of the invasive Cape ivy villain. Stay tuned for the sequel.
In the mean time, if you would like to learn more, see the contact information, links and research articles below.
Mehelis, C.N., Balciunas, J., Reddy, A.M., Van Der Westhuizen, L., Neser, S., Moran, P.J. 2015. Biology and host range of Digitivalva delaireae (Lepidoptera: Glyphipterigidae), a candidate agent for biological control of Cape-ivy (Delairea odorata) in California and Oregon. Environmental Entomology. 44(2):260-276. doi: 10.1093/ee/nvu030.
Acknowledgements:Additional thanks to Dr. Tom Dudley and Dr. Adam Lambert from the UCSB RIVRLAB and Danielle LeFer from California Department of Parks and Recreation for coordinating this momentous day, and to Dr. Patrick Moran, Dr. Scott Portman, Dr. Angelica Reddy, Dr. Chris Mehelis and additional researchers from USDA-ARS in Albany, CA for all of the rigorous testing of this weed and its biological control agents.
One of the reasons I haven’t posted for a bit besides the normal-busy routine is that it is Spring Time! What’s that got to do with anything you ask?
BLOOMS! BLOOMS OF EVERYTHING!
Besides blooms of flowers in the desert (such as those in the Anza-Borrego Desert), we also get blooms of phytoplankton along the coast in Southern California.
Here, in the spring we get very high winds that can result in upwelling events in the coastal ocean, pushing waters offshore and bringing up cold, nutrient rich water from the bottom ocean layers to the top layers.
This increase in nutrients (such as nitrogen, phosphorous, iron, etc.) can result in massive ‘blooms’ or increases in specific phytoplankton species (diatoms, dinoflagellates, etc.), since typically their densities are limited by nutrient availability. During these blooms, whoever wins the space and resource competition will dominate… until they get run down by grazers, parasites or viruses.. or run out of their limiting nutrient. Once these species decline this then provides space/resources for the next dominating species.
These upwelling events also offer AWESOME opportunities for scientists to examine the species dynamics, and the mechanisms that result in some species or functional groups of phytoplankton to dominate over others.
This year, our laboratory (the Caron Laboratory at USC) decided to start our sampling period after we noticed strong winds on April 9th.. and I mean Strong! I was biking to my circus class that evening, and a branch literally flew and hit me.. luckily I was wearing a helmet 🙂 During lab meeting that week, we were all telling each other the horror stories of the strong wind, and realized.. ‘woah!’… we should start our spring sampling asap! So we quickly contacted the amazing Santa Monica Pier Aquarium (Heal The Bay) and received permission to use some of the space there to do our sample processing for three weeks. Then we finalized our schedules, rotating each daily to sample and process the water off of the Santa Monica Pier. Each day at 8:30am, we get to the aquarium, load up our cart with the RBR (an oceanographic instrument that measures temperature, salinity, chlorophyll and dissolved oxygen), a bucket and container for loading up sea water, and a 20 micron plankton net to collect a concentrated water sample. Then by 9am, we are loading up water into our collection container, and then rolling the water back to the aquarium to filter some of it down as fast as possible onto filters that we flash freeze for DNA/RNA extractions. We also preserve some of the whole sea water for relative abundance counts of the different organisms via microscopy, and we sample the water for extraction of chlorophyll and domoic acid (toxin produced by some diatoms). Once we get back to the lab, we inspect the concentrated samples from the plankton net to get a quick overview of who is in the water, and who is the dominating species.
Me on the Santa Monica Pier sampling water
Look how brown the filter is! So much biomass from all the phytoplankton!
Our filter rig to filter the water from the pier
Our set up in the back of the aquarium
This year the sampling has been super interesting! It started off with a diatom and dinoflagellate bloom, and it looks like the diatoms have been CRUSHED by a parasitoid, Cryothecomonas spp.! Once the diatoms crashed, the dinoflagellates increased more, in particular two species are currently dominating: Akashiwo sanguinea and Cochlodinium spp. (species will be determined after we get our molecular sequences back). I also found some tintinnid ciliates parasitized by Eudoboscquella parasitoids.. so beautiful.
Guinardia sp. with Cryothecomonas sp. parasitoid
Dead Guinardia sp with Cryothecomonas sp. parasitoids
Tintinnid (Eutintinnus sp.) with parasitoids (Euduboscquella sp.)
Cochlodinium sp. (dinoflagellate)
Akashiwo sanguinea (Dinoflagellate photo by Jennifer Beatty)
In addition to using molecular sequences for identification of the different taxa, our laboratory also analyzes the RNA sequences (using bioinformatics) to examine gene expression of the different taxa that are increasing and decreasing during the bloom. These methods can help us determine when species are taking up specific nutrients, when they are multiplying, when they are stressed, and even if they are being attacked by parasites. Lastly, my work in particular during this spring bloom will examine the dynamics of these species and their parasites through time using qPCR (quantifies the relative number of the hosts and parasites by comparing samples to standard curves).
We have five more days left of daily sampling, and I will be sure to follow up with another blog on the results of this spring bloom sampling period. I will also post soon about the exciting results from a massive laboratory experiment that I just finished. Stay tuned!
This study was a product of my Delta Science Postdoctoral Fellowship to investigate the mechanisms for effective biological control of the invasive water hyacinth in the Sacramento-San Joaquin River Delta (hereafter “Delta”).
In a nutshell: Two weevils (insects) are currently used all over the world for the biological control of the invasive water hyacinth, including the Sacramento-San Joaquin River Delta, California. They have had variable success, with notable reduction of biomass and cover of this invasive aquatic weed in warmer climates compared to more temperate climates such as the Delta. Although temperature plays a large role in their success, I also investigated the role of genetic variation in the success of these weevils and whether there is lower genetic diversity and heterozygosity in the Delta compared to the native origin of these weevils (South America). To do this, I used polymorphic microsatellite markers (repeating regions of DNA in the genetic blueprints of a species) to detect differences between individuals and between populations. Additionally, as myself and others noticed weevils from the field that appeared to be hybrids of these two species, I examined whether these hybrid-like weevils are genetic hybrids (meaning that they have genetic patterns representative of the genetic blueprints from both species).
In my opinion, the most important findings from this study were:
We found hybrids! This is huge! These two weevils are introduced all over the world for the control of invasive water hyacinth. So now that we know hybridization occurs, it is critical since to understand how hybridization affects their success. For instance, sometimes hybrids can outperform non-hybrids (hybrid-vigor) whereas other times hybridization can decrease performance, as well as population growth (hybrid-breakdown). I am very excited however that Dr. Julie Coetzee’s laboratory in South Africa is now starting to look into the effects of hybridization between these two weevil species.. so stay tuned (I know I will!) .
We found that low genetic variation from demographic bottlenecks (small populations of the weevils being introduced over and over again through the biological control programs), can sometimes be buffered by genetic admixture from multiple introductions. This was one of several findings from this study that was made possible through the unique combination of documented historical records from biological control programs and population genetic analyses, such as those we made with the program, FLOCK.
I also think that the lessons I learned from the process of writing this manuscript were very important, and I detail these below.
Lesson 1: Know when to ask for help
This study culminated out of work that I did at UC Davis, advised by Dr. Ted Grosholz, and in collaboration with researchers, Dr. Paul Pratt and Dr. Kent McCue (USDA/ARS), Dr. Ruth Hufbauer (Colorado State University) and Dr. Pierre Duchesne (Université Laval, Quebec, Canada). The latter two coauthors of whom I actually contacted out of the blue during the analysis and writing portion of the study, since I felt like I needed more guidance from experts in the population genetics and data analysis field. I think knowing when to ask for help is really critical in science (no matter what your academic standing is), and it almost always improves the study to get additional opinions and critique. Think of it as a preliminary peer review before the ultimate peer review!
I also asked several folks that are experts in population genetics for advice on the collection, processing and analysis of the data before and during the start of this project, including: Dr. Jeremy Andersen (UC Berkeley), and Dr. Rick Grosberg and Brenda Cameron (UC Davis) and Dr. Neil Tsutsui (UC Berkeley).
Lesson 2: Be Flexible, and Adapt to let the Data tell the Story
The title of this manuscript felt very suitable to me as ecological data are not always clear-cut, and sometimes it can take some time to wade through the weeds of data and figure out how to tell the accompanying story. This is especially true for when resulting data don’t match up with your original expectations and initial story you thought you would tell. The key to this issue, is don’t try to force your old story on the data… get a second opinion if needed, and be open-minded by letting the data ‘speak’ for itself.
Lesson 3: Work Hard, Be Patient and Persistent
I think with anything that you do, sometimes a final product comes easy… and other times it seems like a long drawn out process. This project fell in the latter category, as it was my first time learning about and implementing a population genetics study, and I was working on the analysis and write-up of this study all while starting a new postdoc in an entirely new study system. I think an important aspect to finishing this project was really persistence. I spent week nights and weekends working diligently on the data analysis and writing and re-writing the paper. I also had to be patient with myself as I had to give myself time to learn the new types of analyses (which means new R packages and code!) and time to read all of the important papers in the study field.
If by chance you are also just starting a population genetic study, and feel a bit lost, please see my three-part tutorial blog posts which hopefully will provide some assistance:
Lesson 4: Implement Self-Deadlines and Advertise them to your CoAuthors
Sometimes its hard to finish something if you don’t have a deadline. So make yourself a deadline, and tell everyone about this deadline, so that you are held accountable for this timeline. I actually had some coauthors that needed me to submit this article to the journal by October 1st in order to meet some of their workplace requirements for publications. Needless to say, I pulled an all-nighter and got it in to the journal by 5am that day.. true story….
Nothing like a little pressure to light up that writing-fire…
Lesson 5: Don’t cut corners
This goes with Lesson 3, on being patient. Towards the end of writing up a big study, you might find yourself just wanting it to be over. You would do anything to not have to think about that project or the data anymore. However, crossing that finish line is actually one of the most crucial components and can make or break your ability to get into a decent journal. Having co-authors often really helps solve this problem, as they will call you out on any cut corners (if they are doing their job), and will suggest critical improvements to the paper that maybe you were thinking about.. but were just initially too lazy to do. Also on this note.. Read the proof-version (final version before being published) of the paper word for word! You don’t want any typos in your finished product.. especially true in your Title, Abstract and Figure Legends!
Lesson 6: Celebrate at Each Stage of Completion
Be sure to acknowledge your accomplishments after you submit the manuscript the first time, after the revisions and acceptance, and after the manuscript goes In Press. After all- you worked hard to get to each of those stages, and celebration will help motivate you for the next time you have to do it all over again!
I am taking a small break from my blog tutorials on using microsatellite markers in population genetic studies to make an exciting announcement: I recently started a new 1-year Post-doc position at the University of Southern California in Dr. Dave Caron’s laboratory (more time pending funding from fellowships)!
Although it is sad that my Delta Science fellowship is over, as it was a wonderful opportunity, I will still be working/writing hard to finish up my publications from this work and I will of course share these with all of you as they are published.
In the mean time- I am moving back into marine study systems to examine the diversity and function of protists in the marine phytoplankton community! Click here to check out the fascinating research in Dave Caron’s lab. In addition to dabbling in several different ongoing projects in Dave’s lab- I am also very excited about starting up some of my own projects (pending funding) on the abundance, diversity and consequences of parasite-host interactions in the phytoplankton community. As some of you might already know- I am an extreme parasite enthusiast, and only recently have researchers started to examine the potential abundance and importance of parasites in the marine phytoplankton community!
Recently, researchers in the Tara-Oceans Expedition found that parasitic interactions were the most abundant pattern in the global marine phytoplankton interactome (Lima-Mendez et al. 2015). Results from the V9-18S tag-sequence processing revealed parasite-host associations that included the copepod parasites: Blastodinium (Dinophyceae: Blastodiniaceae), Ellobiopsis (Marine Alveolate Group I: Ellobiopsidae), and Vampyrophrya (Ciliophora: Oligohymenophorea: Foettingeriida) and alveolate parasitoids of dinoflagellates and ciliates (Lima-Mendez et al. 2015). The alveolate parasitoids in particular were recognized for their top-down effects on zooplankton and microphytoplankton (Lima-Mendez et al. 2015).
Parasitoids are parasites that kill their host in order to complete their development (Lafferty and Kuris 2002) and increased abundance of alveolate parasitoids have been linked to declines of dinoflagellate blooms (Coats et al. 1996, Coats 1999, Chambouvet et al. 2008, Mazzillo 2011, Jephcott et al. 2016) and have been shown to regulate their dinoflagellate host populations in laboratory experiments (Noren 2000, Coats and Park 2002). The most researched alveolate parasitoids include several strains of Amoebophrya ceratii (Marine Alveolate Group II: Syndiniales) . These parasitoids have small flagellated infective stages that penetrate and multiply inside the dinoflagellate host cell, and produce numerous infective flagellates after killing and exiting the host (Cachon & Cachon 1987; Jephcott et al. 2016). For example A. ceratii can produce 60-400 new infective dinospores from its host in less than 48 hours (Chambouvet et al. 2008; Mazzillo 2011), and the generalist parasitoid, Parvilucifera sinerae, can produce 170 to > 6000 zoospores per sporangium, depending on the species and size of its host (Garces et al. 2013), with zoospore release within 72 hours of infecting a host (Alacid et al. 2015).
Below for your viewing pleasure is an example of these parasitoids- the life cycle diagram and life-cycle stages from Alacid et al. 2015, and Alacid et al. 2016 (respectively) of the generalist parasitoid Parvilucifera sinerae, in its host dinoflagellates.
Alacid et al. 2015: Life Cycle of the parasitoid, Parasitoid Parvilucifera sinerae
Alacid et al. 2016: Life cycle stages of Parvilucifera sinerae, a generalist parasitoid, in several of its hosts
So now of course the question you might have is: “why do we need to research these parasites/parasitoids further?” Well, we simply do not know enough about these amazing parasite-host interactions, and most of our knowledge is currently limited to the photic zone of the ocean, and concentrated on just a few of these parasite species (there are many parasites out there just waiting to be discovered!). For those of you that don’t think ‘not knowing enough’ merits more work- my reply to this is that: mortality rates in the phytoplankton community have an incredible significance regarding the total primary production and biogeochemical processes in the ocean. However, how can we account for the mortality rates in the phytoplankton community and consequences for primary production if we are not accounting for a large % of contribution to mortality due to parasites that have not yet been characterized? And this folks.. is the reason why this research should be funded (aside from the obvious fact that parasites are absolutely fascinating, and the evolution and ecology of parasites can tell us a lot about related free-living species as well (that is another blog topic I will save for the future).
Of course my new Post-doc research in this field is still a bit tentative as it depends on gaining further funding- but in the mean time I am posting some lovely photos of parasites (Euduboscquella spp.) in tintinnid ciliate hosts (Eutintinnus spp.) that I have been finding from some local net tows (marine sampling nets that concentrates organisms of different size classes). So exciting- it is like a treasure hunt every time!
Tomont of Euduboscquella parasitoid in Eutintinnus ciliate host
Sporocytes of Euduboscquella in Eutintinnus ciliate host
Sporocytes of Euduboscquella in tintinnid ciliate host, Eutintinnus host
sporocytes of Euduboscquella in Eutintinnus host
References (highly recommended reads also!)
Alacid E, Rene A, Garces E (2015) New insights into the parasitoid Parvilucifera sinerae life cycle: the development and kinetics of infection of a bloom-forming dinoflagellate host. Protist166, 677-699.
Alacid, E., Park, M. G., Turon, M., Petrou, K. & Garces, E. (2016) A game of Russian roulette for a generalist dinoflagellate parasitoid: host susceptibility is the key to success. Front Microbiol 7, 769.
Cachon J, Cachon M (1987) Parasitic dinoflagellates. In: Biology of dinoflagellates, pp. 571-610. Blackwell, New York.
Coats DW (1999) Parasitic life styles of marine dinoflagellates. Journal of Eukaryotic Microbiology46, 402-409.
Coats DW, Adam EJ, Gallegos CL, Hedrick S (1996) Parasitism of photosynthetic dinoflagellates in a shallow subestuary of Chesapeake Bay, USA. Aquatic Microbial Ecology11, 1-9.
Coats DW, Park MG (2002) Parasitism of photosynthetic dinoflagellates by three strains of Amoebophrya (Dinophyta): Parasite survival, infectivity, generation time, and host specificity. Journal of Phycology38, 520-528.
Chambouvet A, Morin P, Marie D, Guillou L (2008) Control of toxic marine dinoflagellate blooms by serial parasitic killers. Science322, 1254-1257.
Garces E, Alacid E, Bravo I, Fraga S, Figueroa RI (2013) Parvilucifera sinerae (Alveolata, Myzozoa) is a generalist parasitoid of dinoflagellates. Protist164, 245-260.
Jephcott TG, Alves-De-Souza C, Gleason FH, et al. (2016) Ecological impacts of parasitic chytrids, syndiniales and perkinsids on populations of marine photosynthetic dinoflagellates. Fungal Ecology19, 47-58.
Lafferty KD, Kuris AM (2002) Trophic strategies, animal diversity and body size. Trends in Ecology & Evolution17, 507-513.
Mazzillo FFM (2011) Novel insights on the dynamics and consequence of harmful algal blooms in the California Current System: from parasites as bloom control agents to human toxin exposure PhD dissertation, University of California, Santa Cruz.
Lima-Mendez G, Faust K, Henry N, et al. (2015) Ocean plankton. Determinants of community structure in the global plankton interactome. Science348, 1262073.