I’ve finally revised the 3D design of pointing and pinning block mentioned in my previous post. This new version of the block has the standard holes of a pinning block (in the center of each level) as well as holes around the outside.
The holes near the edge of each level are for insects glued onto standard points (small paper triangles). By having the holes near the edge, the end of the point with the insect on it can hang over the edge.
This version was designed by me and Julia Amerongen Maddison. I used TinkerCAD for the making the 3D file; TinkerCAD was a joy to use. Some test versions of this were printed with the help of Dan Roach – thank you, Dan! And thanks as well to Pamela Triplett for connecting me with Dan.
For many insects, examination of the genital structures is vital to understand species diversity. The genitalia of each species are often distinct, and species can be easier to distinguish using genitalia than other morphological traits. In carabid beetles, male genitalia tend to have more variation than female genitalia, and so we more often study the former. I’ve shown examples of the differences in male genitalia in my posts about “Morphological subtleties and the value of n>1”, Bembidion subgenusLindrochthus, andBembidion kuprianovii,
When one dissects male genitalia out of a small beetle, one needs a place to store the very small genitalia. A common means of storage is to place it in a drop of glycerol within a very small glass or plastic microvial with a cork or rubber stopper. One can then pin the microvial (through its stopper) on the same pin as the beetle. This has the great advantage that the genitalia is stored with the rest of the beetle, and they are less likely to be disconnected in the future. (One can store the genitalia separately, on microscope slides for example, but that risks their being irretrievably disconnected.)
Storing the genitalia in a microvial under the specimen works well, and allows a great deal of flexibility, but it has several problems. First, it’s a pain to get the very small genitalia of something like a Bembidion out of the microvial, and the very process of taking the genitalia out or putting it back in can lead to damage or loss. Second, every time one wants to compare two specimens, one needs to go through the lengthy process of getting the specimens placed onto microscope slides or the like. Third, given that the specimens are in a fluid, one has to handle them more delicately, and it is harder to compare a large number of specimens at once. Finally, the refractive index of glycerol is far enough away from that of insect cuticle that it is harder to see relevant structures than if the genitalia were in a medium that has a refractive index closer to cuticle.
The genitalia of the carabid beetles I study are relatively flat. We typically study them by looking at their left or right sides, which are the flat sides. Because of this, there’s an alternative to storage in microvials that works quite well: mounting the genitalia in a semi-permanent mounting medium such as Euparal on cards that are pinned with the specimens, as shown in the following picture.
Here’s what the card looks like on its own:
Before I outline how these are made, I should point out that there are other ways to store the genitalia, including embedding them in drop of water-soluble glue on the same point that the beetle is glued to (this was Carl Lindroth’s approach) or by embedding them in a drop of Euparal on a clear sheet of acetate pinned along with the specimen. The approach Carl Lindroth used was not all that much different from placing them in a vial, in that the genitalia need to be extracted from the glue before they can be properly studied. I find placing them in a drop of Euparal on an acetate card also less than ideal, in part as it is more difficult to write a voucher code on the card (which means you can’t easily compare many at once under the scope as you might get them mixed up). There are surely improvements that could be made to the way I make the preps, but I find the method works well enough.
Making genitalic mini-preps
The following instructions presume that you already have dissected out the genitalia, and have cleared it appropriately, and that it is now stored in 100% or 95% ethanol.
Materials you will need
heavy archival paper
a printer (if you are going to print a design)
12 mm circular cover slips. I purchase item number CLS-1760-012 from Chemglass. I typically store them in a clean plastic box, for easy access – it will be important to be able to access them quickly, sometimes with forceps that have a bit of Euparal on them.
Euparal. Euparal can be very fluid and not very viscous when brand new. This is not ideal, as that means it contains a lot of solvent. The ideal Euparal should be much more viscous. I have not fully explored how to make the Euparal more viscous quickly.
100% ethanol (ideally), or 95-96% ethanol
very sharp, clean forceps
Kimwipes or other tissue
a slide warmer with a cover
dissecting microscope with light
flat, long “trays” for holding cards as you process them. You can use microscope slides for this, but I prefer longer, less slippery trays. The ones I use are shown below, and are about 12 cm x 2.5 cm. I made mine out of discarded plastics.
Preparing the cards
If you wish to use a card design, print it on archival, heavy paper. Here’s part of one of the designs I use:
I use Arches hot-pressed, 300 g/m2 (140 lb), watercolor paper. Below is a link to a template that you can use to print on 23 cm x 31 cm (9×12) sheets of the watercolor paper. (Note that when I print this template, I scale the printing to 103%.) I print on a Canon inkjet printer that has archival ink.
Cut the cards out. Then use a standard hole punch to punch out the circles.
Now take each individual card, and turn it over such that the printed side is upmost. This will be side that will have any writing on it, and it is the side that shows the primary view of the genitalia; it will also be the side that is eventually topmost. Using your sharp forceps, dip into the thick Euparal, and get a drop in between the tongs; use that to apply a ring of Euparal around the hole. Use viscous Euparal; thin Euparal will contract too much, forming bubbles, and eventually no longer stick to the cover slip. Even with fairly viscous Euparal, it is a good idea to let the ring of Euparal dry a bit before putting on the cover slip.
Once the Euparal had dried to be very thick, place the coverslip on the card so that is glued over the hole by the Euparal.
Here are some things to watch out for:
Make sure the cover slip doesn’t stick out over one edge of the paper – that will make it more likely the cover slip will eventually come off or break
Don’t put so much Euparal on that it covers the regions of the paper you plan to write on. If you do, the ink you write with may not absorb into the paper, and will be more likely to rub off
Again, use viscous Euparal
Now place the card with cover slip onto your slide warmer. Don’t worry if there is a bit of Euparal that leaks down onto the slide warmer. Leave it there for at least 3 or 4 days, then flip over the card so that the cover slip is against the slide warmer. At this point it will be important that the slide warmer is covered, so that dust doesn’t get into the well of the card. Leave the card on the slide warmer for at least a week (ideally longer) to ensure the Euparal is dry.
In the following figure, you can see the difference between cards whose coverslip was connected with different thicknesses of Euparal. On the left is a card for which thin Euparal was used to connect the coverslip; if you look at it at an angle, with light reflecting off the glass, you can see that most of the space under the cover slip is air, not Euparal; the cover slip will surely just fall right off if it any force is applied to it. The middle card has mostly Euparal connected it to the card; that one should be OK. The one on the right should definitely be good.
With the card completed, if you place it upside down (so that the cover slip is on the bottom), then the card is like a little well slide, with the cover slip forming the bottom of the well, and the sides of the paper hole being the sides of the well. It is into this well that you will place the genitalia.
Adding the genitalia to the card
Once the Euparal on the card is dry, you can add the genitalia to it.
Add the genitalia in standard places and in standard orientations; this is especially important for the parts you will regularly exam. Because the left side of carabid aedeagi is the side most often viewed, I put that side down against the glass. That way, when the preparation is complete, the part we want to view the most will be right up against the glass.
Under the microscope, I extract the genitalia from the ring sclerite, and remove the parameres from the aedeagus in a separate little dish containing 100% ethanol. Once the pieces are all ready, I dip the sharp forceps into a vial of Euparal, and put the drop in the well of the card (which, of course, is now upside down so that the cover slip is against the stage of the microscope). Without cleaning the forceps, I then grab the genitalic parts, ideally all at once, in the forceps (the Euparal on the forceps also helps pick things up), and then put them into the drop of Euparal in the card. I then position all the parts in the well. As you are doing this, consider the following:
Don’t use too big of a drop of Euparal. If you use a lot, then you will not be able to easily position the aedeagus, as it will float around. Use as little as possible so that there is a layer of Euparal over the parts. The parts can stick up; that’s OK.
If you move each piece individually, ethanol will be transferred each time, making the Euparal very liquidy, and potentially dissolving the Euparal that is holding the coverslip onto the paper. This should be avoided
Make sure you use clean forceps.
Filling it with Euparal
Add layers of Euparal, slowly building up the embedding until the well is just full. This may take ten to twenty or more layers. Here are some important things to consider:
As you move slides too and from the slide warmer, clean the sharp forceps. Many surfaces (including those of some slide warmers) will flake off slightly, and those particles can then get into your Euparal and make your preparations dirty.
Make sure you clean off any dust on the surface of the dried Euparal before you add a new layer. You can do this with a clean, small paint brush
Add thin layers. If you add thick layers, then the thicker layers will contain enough solvent to make it more likely the solvent will soften the dried layers the genitalia are in, causing them to float and move.
If the Euparal you are adding is thin, and not viscous (i.e., it contains more solvent), then the layers should be thinner.
If the genitalia are large they may be less likely to float around, and so you may be able to get away with thicker layers.
If the side of the genitalia that is against the glass cover slip is flat, then it is a bit less likely to move around. If the side of the genitalia is rounded, then it can be more likely to move around, and you then need to be even more careful with your layering – make them very thin.
Keep the temperature on the slide warmer low – ideally about 35°C. If it is much hotter than that, the Euparal will become softer, and the genitalia will be more likely to float around.
Place a note next to the cards on the slide warmer saying when the last layer was added.
You want the Euparal to dry between layers, so you will likely want to add new layers no more often than once a week.
Don’t overfill the well. The Euparal in the end should just be flat – not convex. (But note below that sometimes I use much less Euparal – see the next section.)
After adding a layer, put it back onto the slide warmer. Once the well is full (if that is your goal), leave it on the slide warmer for at least a month (I usually leave it for at least 3 months), to ensure the Euparal is very dry.
Note that you can photograph the genitalia well before the preparation is complete.
Adding the second cover slip – or not
Typically I add a second cover slip once the well is full to the brim of dried Euparal. The main point of that second cover slip is to both protect the specimen, but also to make the preparation easier to clean. If dust gets on an open Euparal surface, it can be hard to clean. A clean dry brush can work, but a brush dipped in ethanol usually makes the surface uneven and cloudy, which can be resolved with another layer of Euparal. If the genitalia are really small (e.g., from a 1.5 mm beetle) or very tubular, the danger of their shifting orientation as you add layers is much higher. You may not want to complete the process, and instead simply add enough layers to have a flat surface above the beetle bits, and let it harden, without ever filling it full enough to add the second cover slip.
If you are going to add the second cover slip, you will need to fill up the depression, and you will first need to prepare the cards. Examine the areas around the edges of the hole under the microscope, and and see if there are bumps of dried Euparal or other protuberances that will force the coverslip to be raised off the flat surface. If so, scrape them off as best as you can (this can be done with your sharp forces). You will then need to brush off the surface of the Euparal with a clean, dry brush, to get rid of any dust and lint that has settled there.
Once the surface is clean, you are ready to add the second cover slip. Place a large drop of viscous Euparal onto the preparation (again, I use clean, sharp forceps to do this). You can then pick up the cover slip with forceps, and gently lower it onto the drop (do this at an angle by first having one edge of the cover slip make contact with the paper). If the Euparal is very viscous, enough to cause bubbles to get trapped as you lower the cover slip onto it, then you may wish to dip your forceps into 95-100% ethanol, and touch them to the surface of the Euparal. This will spread a thin layer of ethanol onto the surface, and allow the coverslip to be placed without bubbles. If the Euparal is quite viscous, you may need to push the coverslip down a bit.
You should then look at the preparation from an angle, with light reflecting off the glass (as you did above when you first made the card), and see if Euparal fills the contact area between the paper and the cover slip. If not, add Euparal drops to the edges of the cover slip, and encourage the Euparal to seep under the coverslip. This should also be relatively viscous Euparal.
Check the preparation every few hours for the first day, and add Euparal to the edges of the cover slip as needed. Then check it once a day for at least three days. Once it seems to stabilize, leave it somewhere at room temperature for at least three months before pinning it beneath the specimen.
What to do if it all goes wrong
Sometimes it goes wrong. The most common problem is that the Euparal the genitalia are in softens after a new layer is added, and the genitalia float around and are no longer in the correct orientation. This is frustratingly common, and seems to be usually caused by adding too much Euparal at one time for one or more of the layers. If so, you can simply start again. Place the card in a little dish with 95-100% ethanol, and after a couple of hours, the Euparal will have dissolved, and it will all come apart, and you can do it all again.
In 2006, for George Ball’s 80th birthday, I presented him with a drawing of Dicaelus purpuratus. This was the species that captivated his attention and eventually led to his doing a PhD on the carabid beetle tribe Licinini, of which Dicaelus is a member.
Here’s the final drawing:
Here’s where I did the drawing
I’ve written a post about the drawing method I use, which has varied slightly over the years. For the Dicaelus, I began by enlarging a photograph of the beetle, and printing it on paper.
I then used a soft graphite pencil, and rubbed the back of the paper. That turned it into carbon paper that I could use to transfer some aspects of the photograph onto the Arches Hotpress watercolor paper I use.
I did the transfer by drawing over the photograph using a sharpened 6H pencil. Here’s what was transferred. Note that I have also put a mask over much of the paper.
I then refined the pencil sketch.
The inking started, as well as the colored pencil.
More ink outline, and more colored pencil:
Eventually, once the colored pencil layer was refined, I started adding inks on top of it:
In September of 2009, I arrived in Oregon, excited to begin my new position at Oregon State University. I was also excited to live near Marys Peak, as the top of Marys Peak was the locality of capture of the only known specimen of what was then called Bembidion chintimini. (I’ve since moved that and related species out of the genus Bembidion, and as the genus Lionepha.) A few days after I arrived in Corvallis, I drove up to the top of that mountain to see if I could find the species again. I found some little beetles that might or might not be Bembidion chintimini, but I was a novice with that group of bembidiines, and so I wasn’t sure. I also found a larger, related specimen lower down the slopes of Marys Peak, near Alder Creek Falls, which confused me as well. My efforts to discover the identity of those specimens took a long time, and the threads I was tugging on as I explored caused an unravelling in my understanding of that group of beetles, which I would eventually re-weave into a new tapestry. In the end, this path led to the discovery of a total of four new species, as well as a merging of three others.
In many ways this project was the heart and soul of my first decade in Oregon. I spent many hundreds of hours on this project, traveling and collecting specimens, extracting and sequencing their DNA, making genitalic preparations, studying their structures, photographing them, examining their chromosomes, doing phylogenetic analyses, and, eventually, creating the figures for the paper and writing the text. Perhaps more emotionally compelling was the discovery and solving of many puzzles along the way, especially that revolving around “Bembidion chintimini”. That some of these puzzles were solved by a combination of old-fashioned morphological studies as well as next-generation genomic sequencing of old type specimens, including a 159-year-old LeConte lectotype, made the tale all the more compelling to me.
John Sproul, a former graduate student of mine, helped by sequencing the DNA I extracted from some pinned type specimens (including that LeConte lectotype), as well as by doing important collecting in the Sierra Nevadas of California; for these efforts, he is a co-author of the paper that has finally resulted.
This paper, titled “Species delimitation, classical taxonomy and genome skimming: a review of the ground beetle genus Lionepha (Coleoptera: Carabidae)“, came out recently in the Zoological Journal of the Linnean Society. The paper can be found at https://doi.org/10.1093/zoolinnean/zlz167; if you would like a PDF, email me.
It turns out that I did find a female of “Bembidion chintimini” on Marys Peak that first trip in 2009. It took until the following summer for me to realize that. It took even longer to eventually come to the realization that this species was widespread, and had a much older name (Lionepha erasa LeConte). Here’s a condensed version of the story, as told in the paper:
Investigation of the rarer species, the one here called Lionepha erasa, began in 2010. Dissection of the first recognized males from Marys Peak, Oregon (type locality of Bembidion chintimini) revealed an aedeagus indistinguishable from those from San Juan Island, Washington (type locality of Bembidion lummi). The female holotype of B. chintimini is wingless and has slightly rounded shoulders. However, the Marys Peak population is wing-dimorphic, and winged individuals are in body form no different from the type series of Bembidion lummi. The elytral microsculpture of the holotype of B. chintimini is perfectly isodiametric (against Erwin & Kavanaugh, 1981), thus matching that of B. lummi. Other characters mentioned by Erwin & Kavanaugh as distinguishing the two populations are not consistent with available specimens. The lack of evident morphological differences, combined with effectively identical DNA sequences in specimens from Oregon, British Columbia and Alaska suggested that the Marys Peak populations are the same species as populations further north, and for this reason, Bembidion chintimini and B. lummi were synonymized by Maddison in Kanda et al. (2015).
This left in question the specimens considered to be Bembidion lindrothellus by Erwin & Kavanaugh, which are at first glance similar to the Marys Peak and other populations of ‘Bembidion chintimini’. Specimens classified as Bembidion lindrothellus are reported to be paler, but all specimens mentioned in Erwin & Kavanaugh (1981) are teneral. The unsclerotized aedeagus of the holotype of Bembidion lindrothellus made comparison of internal sac sclerites difficult. However, the internal sac membrane that rests in the left-most position has a species-specific microsculpture in Lionepha, and the microsculpture scales of the holotype of Bembidion lindrothellus from Alaska match those of Marys Peak specimens. A non-teneral male was also collected by Lindroth at the type locality of B. lindrothellus, but was not included in the type series, perhaps as the specimen was housed in Lindroth’s collection in Lund, Sweden. This specimen is presumably the one whose genitalia Lindroth figured as Bembidion brumale (1963: fig. 127f). We have examined that specimen, and it is indistinguishable from specimens of ‘Bembidion chintimini’ from Alaska, British Columbia, Washington and Oregon, including details of the internal sac. Most critically, DNA sequences of the holotype of Bembidion lindrothellus are identical in eight studied genes to those of other specimens from throughout the range (Figs 5–7). It is thus evident that the holotypes of Bembidion chintimini, B. lindrothellus and B. lummi belong to a single species.
However, there is an older name. The type series of Bembidium erasum consists of four females. These specimens have traditionally been considered to belong to the common, widespread species here called Lionepha probata. Females of these two isodiametrically microsculptured species are difficult to tell apart, especially those with less-extreme prothoracic proportions (neither wide nor narrow). Although there are distinctions in the lobe of the female bursa of fully sclerotized individuals, interpretation of tenerals is more tenuous. Specimens in the type series of Bembidium erasum are all teneral, with prothoraces of moderate width, and thus there is no clear morphological evidence to place them to species. The type series was provided by George Suckley (LeConte, 1859), presumably captured during his travels as naturalist for the governor of Washington Territory during 1853–57 (Cooper & Suckley, 1859). The type series is from ‘Oregon’, which at the time encompassed the current area of Oregon, the southern half of what is now Idaho and some parts of Wyoming and Montana (Barry, 1932). Suckley’s travels in Oregon included areas within the range of both species (Cooper & Suckley, 1859), and thus geography provides no clues about species membership. However, DNA data from the lectotype (and two of the paralectotypes; Sproul & Maddison, 2017) makes it clear that these specimens belong to the current species (Figs 5–7; Supporting Information, Fig. S1). Thus, the valid name of this species is Lionepha erasa, with Bembidion chintimini, B. lindrothellus and B. lummi as junior synonyms.
That description of the history does not adequately capture all the many mysteries, proposed and rejected explanations, and confirmed hypotheses along my multi-year path, and the eventual pleasure as the hypotheses became confirmed through the emergence of consistent, repeated patterns. It was one of the more pleasing journeys of discovery I have been on. I’ve previously mentioned a few of the turns and twists in the story, in my posts on Rainy-season beetles and surprises in Lionepha.
In addition to solving the mystery of the small beetles on top of Marys Peak, four new species were discovered along the way and described in the paper:
The larger specimen I discovered on my first trip up Marys Peak belongs to a species that lives along the sides of Alder Creek Falls and on rock seeps in the area, as well as along a creek west of Eugene, Oregon, and in the Trinity Alps of California, is now officially named Lionepha tuulukwa Maddison. I’ve talked about the naming of this beetle here and here.
A species I first became aware of through a single male along Bishop Creek that John Sproul found, is now called Lionepha lindrothi Maddison & Sproul.
The species I found in the central Sierra Nevada of California, and which provided another surprise in the group, is now called Lionepha australerasa Maddison.
A species from western Montana and Wyoming, as well as eastern Oregon and Washington, came to light from specimens collected by my good friend David Kavanaugh. This species is now called Lionepha kavanaughi Maddison.
This paper on Lionepha implicitly tells a love story between me and my beetles. When I look at the final product, I see it as a celebration of the process and fruits of discovery. I also see it as a history of my embracing the Pacific Northwest, and the abundant and diverse life that lives here.
On 16 November 1994, we first made public the prototype version of the Tree of Life Web Project. At that time, it was a series of static web pages, with trees made out of text characters, created by a special version of MacClade.
This was more than three years before Google existed, and more than a year before the research project that led to Google began. In November 1994, Wikipedia was still more than six years away, Facebook was nine years in the future, and YouTube more than 10 years away.
Although the project is still alive (it still gets over a million separate visitors a year, from many countries), it has been relatively dormant, awaiting someone with a passion to take it over and reinvigorate it.
It’s been a good 25 years; it’s hard to believe it’s been that long.
As the ship rolls gently upon the waves,
I look east toward the Tasman Sea,
The relentless surges swallowing time,
As they have for millions of years,
Through turn after turn of the Milky Way,
Beneath and beyond the waves,
The tree of life has grown,
Both struggling for life
Against harsh forces,
And exploding in exuberant replication.
And within our small leaf,
The courage and strength of the principled
Inspires and uplifts,
Giving us strength to seek a brighter future,
And explore this world which has given us birth,
And to which we will return.
There are those amongst us who, through their words and actions, radiate strength, courage, compassion, and generosity, causing a positive ripple to spread across the communities they touch, increasing the good in the world, and casting light against the darkness. George Eugene Ball was such a man.
When I was a teenager, my parents would visit our grandparents, and on the way would drop my brother and me off at the University of Guelph library, where we would happily spend a Saturday or Sunday pouring through the shelves of beetle and spider books, photocopying as many of them as we could. It was in this library that I discovered Ross Arnett’s Beetles of the United States, and, inspired, purchased a copy of that tome. One chapter in particular captured my attention, about Carabidae, written by George E. Ball of the University of Alberta. As a 16-year-old, on 13 December 1974, I decided to reach out to Dr. Ball, in case he might be able to answer my queries about some of the ground beetles I was finding in southern Ontario. I wrote him a four-page letter, but did not expect a response – a famous professor would surely have more important things to do than write a teenager far across the country. To my great surprise and delight, a few weeks later I received an eight-page letter in response, answering my questions in detail, and providing me with the names and addresses of many other carabid systematists to whom I could write.
Decades later, when I wrote a letter telling George how profoundly his letter affected me, and how much it inspired me, and thanking him for it, he replied that the best way to thank him would be to respond similarly when a 16-year-old wrote me asking me questions about her or his favorite beetles.
This typified George. He was a man of principle, duty-bound to serve the causes he believed in, including the people who were part of his community. He would have viewed helping me as a necessity, as he was committed to educating youth and encouraging those who might engage in the honourable cause of discovering and documenting our planet’s biodiversity.
George sought to deflect attention away from himself. He did not want to be viewed as a grand master, separate and above the rest of us. In later years he reluctantly allowed us to celebrate his achievements, not because he wanted to bask in the glory, but rather because it allowed him to see old friends and hear about their discoveries.
George was virtually egoless when it came to science. I remember once sitting with some fellow graduate students as George was explaining some entomological knowledge or systematic principles (I don’t remember which), and at one point he just stopped, looked thoughtful for few seconds, and said, “Wait, why did I just say that? That isn’t true at all”, and he then proceeded to correct himself. This was a profound message to those of us concerned about social standing: it told us that science required us to focus on honesty and the pursuit of knowledge, even if that meant proving ourselves wrong in public. We learned that science was not about us; our pursuit was bigger than any of us.
One of George’s most important traits was his positivity toward others. He was almost always generous regarding others, and emphasized their better traits. He would not say anything negative about anyone in frustration or in spite, an inspiring demonstration for those of us not always so kind. Even in situations where it would be better to be honest about the negative traits about someone, he struggled to express them. To be at fault in this direction is vastly more preferable than the alternative.
One might think a man such as this would set such a daunting example that he would not inspire others, as he would be viewed as so distinct from the rest of us that we could not possibly follow his path. But George was humble, and that gave us a sense that we, too, could live by those principles. He was also not perfect, and did not hide his imperfections, and through those we could see his struggle, and what he had achieved by force of will; this led us to believe that we too could achieve our goals with honesty and hard work.
You will be missed, George, but the ripple of goodness and light that you created will continue to spread and amplify.
Yesterday, after about three years in gestation, we released Zephyr 2.0. Zephyr is a Mesquite package that manages interactions with phylogeny inference packages including RAxML, GARLI, PAUP, and TNT.
The most notable additions to Zephyr 2 include implementation of the SOWH test, CIPRes support, much better interapplication communication, and more extensive support for PAUP. Many bugs were also fixed, and other improvements made.
The Swofford-Olsen-Waddell-Hillis test allows one to test particular aspects of phylogenetic structure, such as the presence of a hypothesized clade. Given a Mesquite file containing:
the data matrix
a constraint tree showing just the phylogenetic structure to be tested (e.g., a tree showing just the one clade with everything else as polytomies)
the model tree: the best tree with branch lengths that fits that constraint (inferred in a constrained analysis)
models of evolution (e.g., GTR+I+G) with parameter values inferred from the matrix,
then the SOWH feature in Mesquite will automatically find the observed value of the test statistic using whatever tree inference program you choose among those Zephyr supports, and simulate data many times on the model tree, calculating the test statistic for each simulated matrix. It will show you the p-value as the analyses goes along, and gives a report once you have decided you have done enough replicates.
In the figure above, the model tree on which data are simulated is shown in the middle, with the results from the SOWH test on the right. (In this example, only four replicates were conducted; to get an accurate estimation of the p-value, many more would need to be done.)
CIPRes (CyberInfrastructure for Phylogenetic Research, http://phylo.org) provides a gateway for doing phylogenetic inference on a fast cluster of computers. Zephyr 2 allows one to run analyses on CIPRes from within a Mesquite session, and will harvest the results once done and move the trees into the Mesquite file.
Zephyr 2 has many improvements under the hood, including much better communication mechanisms between Mesquite and the external program. Among the improvements are the option to have Mesquite directly start the external program (as opposed to asking the operating system’s shell to do that), which gives Mesquite more control over the process.
Better PAUP support
Zephyr 2 now provides a means to do likelihood, distance, and SVD quartets analyses using PAUP from within Mesquite.
I started learning how to do stained glass pieces over the summer, and for my first piece I decided to honor our Discovering Insect Species class by doing the head of an undescribed species of Bembidion (Trepanedoris) that we discovered. I finished the piece on Tuesday; here it is: