biology articles imaging

Mitotic waves and gastrulation in a fly embryo

Last year I published a snapshot of a mitotic wave in a fruit fly embryo. Here’s the video of that same embryo going through cleavage (nuclei divisions) and gastrulation (cell movements):

Mitotic waves (cycles of nuclei divisions) and gastrulation in the fruit fly Drosophila melanogaster. Also on Wikimedia Commons.

Mitotic waves

What you see at the beginning of the movie are the cycles of synchronous nuclei divisions. They happen in waves from the posterior to the anterior side (from right to left). Fly embryos undergo fourteen cleavage cycles after fertilization, but the movie only starts on the tenth. After each cycle, the embryo gets more packed with nuclei until they cover its entire surface.

Foe and Odell 1989 Nuclear cycles
Cycles of nuclear divisions in early Drosophila embryogenesis (Foe and Odell 1989).

At this point, the embryo is still a syncytium, that is, a single cell with many nuclei (yeah, that’s how flies do it). But on the 14th division cycle, about 11s into the movie, the embryo cellularizes, each nuclei being encapsulated by cell membranes. It’s a curious process, though not visible in this video.


Once cells form, the embryo begins to gastrulate. Gastrulation in flies is complicated, and many events happen at the same time. Cells move, invaginate, disappear, flow around the embryo, and start dividing again, again, and again. This movie shows about 3 hours of development, but embryogenesis continues for 24 hours post-fertilization until the embryo hatches out of the egg as a larva.

Foe and Odell 1989 Gastrulation
Developmental events during the gastrulation of Drosophila (Foe and Odell 1989).


The fly in the video is a transgenic line containing two types of fluorescent proteins. One is a green fluorescent protein (GFP) which is attached to a histone (H2A), a protein that binds to DNA in the cell nucleus. The other is a variant red fluorescent protein known as mCherry, which is fused to a protein that attaches to cell membranes (GAP43). In this way, we can see where the chromatin and cell membranes are located in the embryo during development (and learn something from it).


After collecting embryos, I glued them sideways on a tiny glass coverslip attached to the sample holder of the microscope, a Zeiss Z.1 Lightsheet. The scope has lasers to excite the fluorescent proteins which in turn emit light at different wavelengths. Filters allow us to capture the individual signals simultaneously. This is crucial for fast acquisition and makes two-color time-lapses possible.

I set the microscope to acquire 30 slices spaced by 3 µm, from the outer surface of the left side to the middle section of the body. One timepoint was taken every 35 s.


After the recording, I performed some image processing steps using Fiji/ImageJ. While 3D renderings for this type of data look great, I decided to flatten the image to 2D.

First, I ran Despeckle to reduce some noise (it’s low but helps smoothen the image). Second, I ran Subtract background... (with rolling=20) to remove out-of-focus information. Fly embryos have a dense yolk sac inside that can blur the fluorescence. Finally, I did a maximum intensity projection using the Z Project... command. This looks through the 30 slices in each pixel position, and only keeps the maxima values. It’s a practical way to visualize 3D data in 2D.

Lastly, we can color-code each fluorescent marker differently for better visualization. Some colors fit well together, others don’t. To map pixel intensities to color gradients in ImageJ we use lookup tables (or LUTs). Here, I chose a purple-yellow gradient named mpl-inferno for the DNA signal, a perceptually uniform colormap I like a lot, and a standard grayscale for the membranes (gray is great).

The end

That’s it, I hope you enjoy the video! I’ve uploaded a copy to Wikimedia Commons for eternity. Feel free to use it and if you need additional info, please contact me.

imaging articles code

ImageJ macro to synchronize and combine image stacks

The embryos I study rarely develop in perfect synchrony. That means that when I film them under the microscope some embryos will be younger—or older—than others.

ImageJ macro with Drosophila embryo
Using an ImageJ macro to help me analyze movies of Drosophila embryos.

For this reason, I often need to synchronize the recordings to make sure they all begin at the same embryonic stage. When the movies are synchronized I can combine them side-by-side, and it becomes much easier to compare and spot differences between two embryos.

ImageJ macros save time

Combining movies in Fiji/ImageJ is straightforward using the Combine... command. But synchronizing is way harder. It depends on human classification and involves some calculations and stack juggling that can (and will) become tedious.

To help me out, I wrote a small ImageJ macro available here: SyncAndCombineStacks.ijm. Follow below to see how it works.

Combined movies without syncing

That’s what unsynchronized movies look like. I combined them fresh off the microscope without any synchronization:

Two embryos of the fruit fly Drosophila melanogaster. Both were acquired in the same microscopy session. The top embryo is older than the bottom embryo.

Combined movies after syncing

Here are the same two movies now synchronized by the embryonic stage:

The same two embryos are now synchronized.

How it works

The macro performs the hard work. It calculates how many frames to trim from each stack. Then it duplicates the selected range of frames common to both stacks. Finally, it combines the synchronized recordings into a single image stack. All you need to do is to select the corresponding frames between the two stacks.

Step-by-step instructions

Here are the instructions step-by-step:

  1. Open both image stacks in ImageJ.
  2. Adjust the contrast if needed (before running the macro).
  3. Select a reference frame in the top stack (e.g. stage easy to recognize).
  4. Select the correspondent frame in the bottom stack.
  5. Run the macro and fill in the dialog parameters.
  6. Click OK, wait a few seconds, and check if the synchronization is good. Otherwise, re-run with different parameters.


I’ve also recorded a small screencast:

Note! The macro does not touch the original stacks, but it outputs an RGB Color stack. There are a couple of reasons for that. Converting to RGB avoids contrast issues when the stacks have different pixel intensities. It also prevents quirks in video players that can’t handle 16-bit movies. But if you need to perform image analyses on the final stack, remove this option. I may add a checkbox for that in the future.

articles code

Convert video to animated GIF

Something that I began doing more often is converting videos of developing embryos or marine invertebrates to animated GIFs. But how to do this conversion without affecting the quality of the video?

A jellyfish moving its tentacles. Source: Cifonauta.

Some time ago I found this guide to convert videos to high-quality animated GIFs using the tool FFmpeg. The trick is to generate a color palette based on the original video to improve the color quality of the GIF. Based on this guide I created a small bash script to make my life easier and perhaps yours too ;)

Check it in

music articles

The Pluteus Trip

The Pluteus Trip is a music compilation that I created inspired by the life of these nifty echinoderm larvae named pluteus. It was released more than ten years ago in my (now defunct) music blog ccNeLaS.

The album is freely available at:

Please find the original description below and enjoy the trip!

The Pluteus Trip front cover.
Front cover of The Pluteus Trip.

Plutei are born in the seawater. They represent a specific life stage (larva) of some marine invertebrates, the Echinoderms. Most of them are less than 1mm long, so tiny that inertial forces are dominated by viscous forces of the water.

Just imagine if air was honey and we had to go for a walk… Plutei can swim and feed in this environment using their long arms and cilia. However, Plutei are ephemeral. They swim (and eat) for weeks or maybe months, before something else takes place.

Currents can take them really far away from the place they were born. Millions of Plutei are born at once. How many would survive? How many would be thousands of miles away? How many would get proper food and not be eaten?

Plutei carry the tissue of adults inside them. The food they eat goes to adult tissues. In the end, the adult in formation takes over the larval body and the Pluteus is gone.

Plutei are part of the ocean’s hidden life. Organisms we can’t see easily, but that certainly got in between our toes when walking along the beach, or were swallowed during a swim…

The Pluteus Trip back cover.
Back cover of The Pluteus Trip with the song list.
biology articles

Is Lineus longissimus the longest animal on Earth?

Which is the longest animal on this planet?

Last year the PeerJ journal published an article about the largest marine animals. The neat infographic accompanying their tweet immediately got my attention (if the figure is not showing up, check it in full resolution here):

For a couple of milliseconds, I thought the scale bar below the whales was a worm. Why? Because here in Norway, at the Sars Centre, we study a ribbon worm named Lineus longissimus – the longest animal on Earth.

Bootlace worm Lineus longissimus
The bootlace worm Lineus longissimus is a nemertean (=ribbon worm) known for its long body length. Photo: Wikimedia Commons

Or at least it’s supposed to be. That is what’s in the Guinness World Records (Cawardine 1995), in Wikipedia, in books and papers, and recently on the BBC. Every year on Bergen’s Research Day we tell the kids that this thin dark-brownish worm they are looking under the scope can reach 60 meters long! But how do we know?

Finding Lineus longissimus

Me (1.90m) with one L. longissimus specimen in the animal facility. Longest animal on Earth? Photo: Anlaug Boddington
Me (1.90m) holding one Lineus longissimus specimen in the animal facility. Photo: Anlaug Boddington

We collect live specimens of Lineus longissimus by dredging the bottom of the Norwegian fjords near Bergen. They are active, bear a good dose of charisma and survive well in the laboratory. They are also voracious predators and like to feed on annelids, which are usually swallowed whole:

But so far, the longest individual we’ve found is near two meters long. How could it reach sixty?

Digging up old literature

Here is what the most-cited description says:

The longest known worm is the bootlace worm (Lineus longissimus), a kind of ribbon worm or nemertine (Nemertea) found in the shallow waters of the North Sea. A specimen which washed ashore after a severe storm at St. Andrews, Fife, UK, in 1864, measured more than 55 m (180 ft) in length.

Cawardine 1995

In 1864, a near-sixty meters worm was found in Scotland. Found by who? Measured how? Preserved where? Who wrote the original report? All the links mentioned above converge to one source: Cawardine (1995). But to my surprise, his book does not cite the original report. For this reason I decided to check out what humanity knows about the length of Lineus longissimus.

I knew the date. The location suggested that the report must be in some British literature. To find the original sixty-meter observation I resorted to the great Biodiversity Heritage Library (BHL), which helped me several times to access the classical literature during my doctorate work. BHL has a neat feature where you can search the literature by a species name and the bootlace worm Lineus longissimus is present in many publications.

Here is an almost exhaustive compilation of reports with the length of Lineus longissimus:

ReferenceMaximum lengthComment
Sowerby (1806)Many fathomThis is estimated length, not a direct measurement (1 fathom = 1.83 meters).
Pennant (1812)9.1 m
Davies (1816)28 mDirect observation: 6.7 meters after fixation. At least four times this size when alive. Estimated between 22–28 meters (12–15 fathom).
Schweigger (1820)1.2–4.5 m
Edwards (1846)27 m
Newman (1848)27 m
Thompson (1849)4.0 mDirect observation: 1 meter after fixation. Estimated 4 meters (12 feet) when alive.
Thompson (1856)3.7 m
Leuckart (1859)0.5 m
Johnston (1865)4.3 mDirect observation.
McIntosh (1873–1874)4.5–9.1 mEstimated (15-30 feet).
Claus (1876)1.5 m
Carus (1885)4.5–13.7 m
Knauer (1887)2.0 m
Lacaze-Duthiers (1890)7.0 m
Claus (1891)4.6 m
Oudemans (1892)12 m
Feuille des jeunes naturalistes (1893–1894)25 m
Liverpool Biological Society (1893–1894)0.6–0.9 m
Bürger (1895)11 m
Bürger (1895)27 m
Bürger (1895)30 m
Duncan (1896)4.3 m
Haeckel (1896)12 m
Société zoologique de France (1896)1.0 m
Page (1906)6.1 m
Schmidt (1912)15 m
Société de biologie (1914)2.5 m
Thomson (1916)25 m
Brehm (1918)30 m
Boulenger (1936)27 m
Wieman (1938)30 m
Field Museum of Natural History bulletin (1977)55 mCites St. Andrews specimen as length estimated to 180 feet.
Academia de Ciencias de Cuba (1994)30 m

What I found is that nobody has ever captured (and reported) a live Lineus longissimus more than 10 meters long (but see below!). The longest length is reported by Davies (1816) as 28 meters, and this is an approximation based on how the animal shrinks upon fixation. All subsequent reports simply reproduce this number as a maximum length.

What about the sixty meter worm from St. Andrews?

An ocean’s giant

I initially missed it and months went by… until Jon Noremburg gave the answer in a comment to the above necrophagy video: the St. Andrews report comes from McIntosh.

This is unquestionably the giant of the race, and even now I am not quite satisfied about the limit of its growth, for after a severe storm in the spring of 1864 a specimen was thrown on shore at St. Andrews which half filled a dissecting jar eight inches wide and five inches deep. Thirty yards were measured without rupture, and yet the mass was not half uncoiled.

McIntosh (1873–1874: pg. 183)

Thirty yards (=90 feet or 27 meters) is the measured length of half-worm. Therefore, a whole-worm measures the double, 60 yards (=180 feet or 55 meters). Right? Well… if the worm was partly in knots, how do we know that the measured part is actually half of the length? We don’t.

Calculating the volume of a worm

But the report has a crucial piece of information. The worm was said to fill half of a 8 by 5 inches jar. With some basic maths we can calculate the volume of the worm and estimate its length from that. Assuming that the jar was cylindrical:

V = \pi r^2 h

Radius (r) is 4 inches and the height is 2.5 inches (half of the 5 inches jar). Thus, the volume of the worm equals:

V = \pi \times 4^2 \times 2.5

Or converting to meters:

V = \pi \times 0.102^2 \times 0.064 = 0.0020918483179487 m^3

We can then assume the worm is a cylinder and estimate its height for any given diameter. The height will be length of the worm.

h = \frac{V}{\pi r^2}

The width of L. longissimus ranges from 2–10 mm. I used the formula above to calculate the estimated lengths of the famous St. Andrews worm:

Width (millimeters)Length (meters)

The most conservative estimate (10 mm of diameter) results in an almost 30 m long worm, suggesting that the St. Andrews specimen might indeed have been over 30 m! How much longer is hard to say.

The specimens we have in the lab are between 1 and 4 mm wide, but most are below or near a meter long. It would be interesting to know how the body width scales with the body length in Lineus longissimus.

A specimen of L. longissimus over a ruler.
A specimen of L. longissimus chilling over a ruler.

A worm as long as a blue whale

In any case, the St. Andrews specimen was at least +30 meters, which is a comparable size to the longest ocean giants  the Lion’s Mane Jellyfish (36.6 m) and the Blue Whale (33 m) (McClain et al. 2015). In favor of the nemertean, the volume estimation seems to corroborate his majestic length.

McClain and collaborators (2015) stress the difficulty of estimating the size of large marine animals. Lack of data, biased sampling or simply feasibly measuring a 30 m animal complicates body size assessment. Nemerteans have an additional aggravating factor: they shorten or elongate with ease. Possibly, volume is a more accurate measure of body size for these slim worms.

The authors also highlight that the greatest reported size is quite different from the mean population size. This finding seems true for Lineus longissimus as well. Despite the largest 30–55 m estimate, most of the reports describe lengths not longer than 10 m, often ranging from 1–5 m. Which might show best the normal size distribution of Lineus longissimus populations.

The longest animal on Earth?

Sixty meters might be far-fetched, but there is relatively good evidence that this nemertean can reach – and maybe surpass – the impressive length of a blue whale, thus placing Lineus longissimus indeed within the world’s ocean giants.


Carwardine, M. (1995). The Guinness Book of Animal Records. Guinness World Records Limited.

McClain, C.R., Balk, M.A., Benfield, M.C., Branch, T.A., Chen, C., Cosgrove, J., Dove, A.D.M., Gaskins, L.C., Helm, R.R., Hochberg, F.G., Lee, F.B., Marshall, A., McMurray, S.E., Schanche, C., Stone, S.N. & Thaler, A.D. (2015). Sizing ocean giants: patterns of intraspecific size variation in marine megafauna. PeerJ 3, e715.

biology articles

Segmentation, a question of boundaries

Read the previous section: Spiral cleavage, an oblique matter.

Annelids, arthropods and vertebrates show a remarkable morphological diversity (Chipman, 2010). Beneath this multiplicity of shapes and forms lies a common pattern of body organization—a trunk divided into repeated parts. This pattern and the developmental process that generates it are known as segmentation (Minelli and Fusco, 2004). While the vertebrate trunk is divided into somites1 (a portion of the mesoderm), the body of annelids and arthropods is divided into intricate repeated compartments spanning the ectoderm and mesoderm—the segments (Scholtz, 2002). The morphological similarity between these body segments previously was taken as support for a kinship between Annelida and Arthropoda, in a group called Articulata (Scholtz, 2002; Seaver, 2003). In this scenario, segmentation would have evolved only once in the protostomes and once in the deuterostomes (Davis and Patel, 1999; Peel and Akam, 2003; Seaver, 2003).

Taxa with a segmented trunk. Annelida: the holoplanktonik polychaete Tomopteris sp., Arthropoda: a mantis shrimp (Stomatopoda), Vertebrata: a Teleostei fish larva. Yellow lines mark the anterior and posterior boundary of one segment. Image on the right is a closeup of the ectodermal segmentation of the fire worm Eurythoe complanata. Images not to scale. Photos by Alvaro E. Migotto (Migotto and Vellutini, 2011).
Taxa with a segmented trunk. Annelida: the holoplanktonik polychaete Tomopteris sp., Arthropoda: a mantis shrimp (Stomatopoda), Vertebrata: a Teleostei fish larva. Yellow lines mark the anterior and posterior boundary of one segment. Image on the right is a closeup of the ectodermal segmentation of the fire worm Eurythoe complanata. Images not to scale. Photos by Alvaro E. Migotto (Migotto and Vellutini, 2011).

Analyses arising from the area of molecular phylogenetics have disputed the monophyly of Articulata, suggesting that annelids and arthropods occupy different branches of protostomes, the Lophotrochozoa (=Spiralia) and Ecdysozoa, respectively (Aguinaldo et al., 1997; Eernisse, 1998). This phylogenetic hypothesis indicates that annelids and arthropods are more closely related to groups without body segmentation than to each other (Seaver, 2003); a topology that favors the independent evolution of annelid and arthropod body segmentation, in addition to the independent evolution of the different segmented tissues of vertebrates (Graham et al., 2014). Subsequent phylogenetic studies continue to corroborate the distant relationship between annelids, arthropods and vertebrates (Dunn et al., 2008; Dunn et al., 2014; Edgecombe et al., 2011; Hejnol et al., 2009), reinforcing the homoplasy of their body segmentation.

Remarkably, the molecular mechanisms of body segmentation in arthropods and vertebrates show a number of striking similarities (Damen, 2007; Davis and Patel, 1999; Kimmel, 1996; Patel, 2003; Peel and Akam, 2003; Seaver, 2003; Tautz, 2004). These molecular similarities were taken as evidence to support the homology of bilaterian segmentation (De Robertis, 1997; De Robertis, 2008; Dray et al., 2010; Kimmel, 1996), despite the opposing data from phylogenetics. To reconcile this apparent conflict between developmental and phylogenetic data, we must apply a comprehensive evolutionary approach to the problem.

The concept of segmentation is often used in a typological—and not evolutionary—manner (Budd, 2001). The result is a taxonomic bias, where the evolution of segmentation is regarded from the point of view of the groups considered to be segmented, i.e., annelids, arthropods and vertebrates (Budd, 2001). As a matter of fact, there is no conceptual basis to restrict segmentation to these three groups, because the repetition of parts along the body axis (Budd, 2001; Hannibal and Patel, 2013; Minelli and Fusco, 2004) also occurs in varying degrees in other bilaterians—usually considered to be pseudosegmented or unsegmented (Budd, 2001; Minelli and Fusco, 2004; Scholtz, 2002; Willmer, 1990).

Another aspect to be considered is that segmentation—as much as spiral cleavage—is a complex of characters that ought to be individually compared between taxa (Scholtz, 2010). Breaking down segmentation into comparable traits (Scholtz, 2010), such as seriated nerve chords, segmented mesoderm or ectodermal boundaries, should provide a better overview of their evolutionary history.

Nevertheless, the sole comparison of traits between distantly related groups can still be misleading for understanding the evolution of a character (e.g., trunk segmentation), because the ancestral conditions of closer taxa are unknown. Since developmental mechanisms can be coopted to nonhomologous structures (Shubin et al., 2009), the phylogenetic context of a character is essential to distinguish homology from convergence. A recurrent proposal to better understand the evolution of segmentation is to expand taxonomic sampling (Arthur et al., 1999; Budd, 2001; Couso, 2009; Davis and Patel, 1999; Minelli and Fusco, 2004; Patel, 2003; Peel and Akam, 2003; Seaver, 2003; Tautz, 2004). Thus, examining segmentation traits in a wider range of taxa, including those without obvious segmented features, might help us to grasp the evolution of the developmental mechanisms that form repeated body parts in bilaterians.

This text is the final section of my PhD thesis (published on this blog).


Aguinaldo, A.M. et al., 1997. Evidence for a clade of nematodes, arthropods and other moulting animals. Nature, 387(6632), pp.489–493. Available at:

Arthur, W., Jowett, T. & Panchen, A., 1999. Segments, limbs, homology, and co-option. Evolution & development, 1, pp.74–76. Available at:

Budd, G.E., 2001. Why are arthropods segmented? Evolution & development, 3(5), pp.332–342. Available at:

Chipman, A.D., 2010. Parallel evolution of segmentation by co-option of ancestral gene regulatory networks. BioEssays: news and reviews in molecular, cellular and developmental biology, 32(1), pp.60–70. Available at:

Couso, J.P., 2009. Segmentation, metamerism and the Cambrian explosion. The International journal of developmental biology, 53(8-10), pp.1305–1316. Available at:

Damen, W.G.M., 2007. Evolutionary conservation and divergence of the segmentation process in arthropods. Developmental dynamics: an official publication of the American Association of Anatomists, 236(6), pp.1379–1391. Available at:

Davis, G.K. & Patel, N.H., 1999. The origin and evolution of segmentation. Trends in cell biology, 9(12), pp.M68–72. Available at:

De Robertis, E.M., 1997. Evolutionary biology. The ancestry of segmentation. Nature, 387(6628), pp.25–26. Available at:

De Robertis, E.M., 2008. The molecular ancestry of segmentation mechanisms. Proceedings of the National Academy of Sciences of the United States of America, 105(43), pp.16411–16412. Available at:

Dray, N. et al., 2010. Hedgehog signaling regulates segment formation in the annelid Platynereis. Science, 329(5989), pp.339–342. Available at:

Dunn, C.W. et al., 2008. Broad phylogenomic sampling improves resolution of the animal tree of life. Nature, 452(7188), pp.745–749. Available at:

Dunn, C.W. et al., 2014. Animal Phylogeny and Its Evolutionary Implications. Annual review of ecology, evolution, and systematics, 45(1), pp.371–395. Available at:

Edgecombe, G.D. et al., 2011. Higher-level metazoan relationships: recent progress and remaining questions. Organisms, diversity & evolution, 11(2), pp.151–172. Available at:

Eernisse, D.J., 1998. Arthropod and annelid relationships re-examined. In Arthropod Relationships. The Systematics Association Special Volume Series. Springer Netherlands, pp. 43–56. Available at:

Graham, A. et al., 2014. What can vertebrates tell us about segmentation? EvoDevo, 5(1), p.24. Available at:

Hannibal, R.L. & Patel, N.H., 2013. What is a segment? EvoDevo, 4(1), p.35. Available at:

Hejnol, A. et al., 2009. Assessing the root of bilaterian animals with scalable phylogenomic methods. Proceedings. Biological sciences / The Royal Society, 276(1677), pp.4261–4270. Available at:

Kimmel, C.B., 1996. Was Urbilateria segmented? Trends in genetics: TIG, 12(9), pp.329–331. Available at:

Migotto, A.E. & Vellutini, B.C., 2011. Cifonauta – marine biology image database. Cifonauta, an image database for marine biology. Available at: [Accessed December 16, 2015].

Minelli, A. & Fusco, G., 2004. Evo-devo perspectives on segmentation: model organisms, and beyond. Trends in ecology & evolution, 19(8), pp.423–429. Available at:

Patel, N.H., 2003. The ancestry of segmentation. Developmental cell, 5(1), pp.2–4. Available at:

Peel, A. & Akam, M., 2003. Evolution of segmentation: rolling back the clock. Current biology: CB, 13(18), pp.R708–10. Available at:

Scholtz, G., 2002. The Articulata hypothesis – or what is a segment? Organisms, diversity & evolution, 2(November 2001), pp.197–215. Available at:

Scholtz, G., 2010. Deconstructing morphology. Acta zoologica , 91(1), pp.44–63. Available at:

Seaver, E.C., 2003. Segmentation: mono- or polyphyletic? The International journal of developmental biology, 47(7-8), pp.583–595. Available at:

Shubin, N., Tabin, C. & Carroll, S.B., 2009. Deep homology and the origins of evolutionary novelty. Nature, 457(7231), pp.818–823. Available at:

Tautz, D., 2004. Segmentation. Developmental cell, 7(3), pp.301–312. Available at:

Willmer, P., 1990. Body divisions – metamerism and segmentation. In Invertebrate Relationships: Patterns in Animal Evolution. Cambridge University Press, pp. 39–45. Available at:

  1. In addition to the somites, vertebrates also show segmentation in the rhombomeres and in the pharyngeal archs; segmented structures that likely evolved independently in the deuterostome lineage (Graham et al., 2014).
articles biology

Spiral cleavage, an oblique matter

Read the previous section: Larvae as the epitome of evolution.

By the end of the 19th century, a series of biologists had dedicated themselves to following and discovering the fate of individual cells of an embryo during ontogeny. These works, known as cell lineage studies1, were critical to disambiguate the relationship between ontogeny and phylogeny, directly challenging the idea of recapitulation (Guralnick, 2002; Maienschein, 1978).

The detailed work of the cell lineage biologists Edmund B. Wilson, Edwin G. Conklin, Frank R. Lillie and others, revealed something remarkable. After carefully tracing the embryonic cells of different organisms, they discovered that animals such as molluscs, annelids, nemerteans and polyclad flatworms, whose adult stages are so different, actually share a similar embryogenesis2 (Child, 1900; Conklin, 1897; Heath, 1899; Lillie, 1895; Mead, 1897; Wilson, 1892). Their embryos show the same cleavage pattern, in which cell divisions occur with the mitotic spindles oblique to the animal/vegetal axis, switching direction (clockwise and counterclockwise) at each division cycle (Costello and Henley, 1976; Hejnol, 2010; Henry and Martindale, 1999; Lambert, 2010). A quartet of vegetal macromeres sequentially gives rise to animal micromeres, and the resulting symmetry of these cleaving blastomeres, when viewed from the animal pole, was described as spiral. This developmental pattern thus became known as spiral cleavage (Wilson, 1892).

The spiral cleavage pattern.
The spiral cleavage pattern. (A) Animal pole view of a generalized spiral-cleaving embryo. Arrows indicate the direction of cell divisions. Developmental sequence based on (Conklin, 1897). (B) Schematic diagram of cell divisions in the D quadrant in a lateral view (top: animal pole, bottom: vegetal pole). Cells are named with the standard spiral cleavage notation(Child, 1900; Conklin, 1897; Wilson, 1892). Representation based on Lambert (2010).

Because the cell divisions are stereotypic, individual blastomeres can be followed and compared between spiral-cleaving taxa in a fairly consistent manner. The ability to compare blastomere fates at this unprecedented cellular-resolution uncovered a surprising similarity in the fate maps of spiral-cleaving embryos (=annelids, molluscs, nemerteans and polyclad flatworms). The iconic example being the 4d mesentoblast, a well-conserved mesoderm precursor (Lambert, 2008). Overall, despite having the oblique cell divisions as an idiosyncrasy, spiral cleavage is understood today as a complex of developmental characters (Costello and Henley, 1976; Hejnol, 2010; Henry and Martindale, 1999; Lambert, 2010).

The empirical findings of cell lineage studies raised several important evolutionary questions regarding the evolution of development and the establishment of homologies (Guralnick, 2002). What are the underlying causes behind embryonic cleavage patterns—mechanical forces acting on the embryo or inherited historical factors? Are the events of early development necessary to build the adult characters? Is there an embryological criterion for homology? The ideas progressively moved towards a more evolutionary view of development, where ontogeny is not “a brief and rapid recapitulation of phylogeny” but an inherited product of evolution and subject to modification (Guralnick, 2002).

Even though most cell lineage biologists initially denied the systematic value of embryonic cleavage patterns, mainly in opposition to recapitulation (Guralnick, 2002), it was difficult to argue against the striking similarity between spiral-cleaving embryos, and dismiss their potential kinship3. Schleip (1929) was the first to propose a group to contain the animals displaying spiral cleavage—the Spiralia.

Recent metazoan-wide phylogenetic analyses corroborate the kinship between spiral-cleaving taxa, in a major protostome clade that is sister to the Ecdysozoa (e.g., insects) (Dunn et al., 2014). The latest works in protostome phylogenomics (Laumer et al., 2015; Struck et al., 2014) suggest that Spiralia (=Lophotrochozoa in some cases, see Hejnol (2010)) contains not only the typical spiral-cleaving groups, but several other taxa. Some spiralians (=animals that belong to the clade Spiralia) do not show any clear trace of spiral cleavage, such as bryozoans, brachiopods, gastrotrichs and rotifers, while others do exhibit spiral-like characters, such as gnathostomulids (Riedl, 1969), phoronids (Pennerstorfer and Scholtz, 2012) and entoprocts (Marcus, 1939; Merkel et al., 2012). What can we say about the evolution of these disparate cleavage patterns?

The spiral arrangement of embryonic blastomeres is present in the three main clades of Spiralia (Gnathifera, Lophotrochozoa and Rouphozoa), suggesting that this character is ancestral at least to the Lophotrochozoa-Rouphozoa clade. This implies the spiral cleavage pattern was lost during the evolution of gastrotrichs, brachiopods, bryozoans and maybe rotifers. How did these groups lose spiral cleavage? Which aspects of a typical spiral-cleaving embryo did they lose, in addition to the spiral arrangement of the blastomeres? Are there any remnants of spiral cleavage?

The comparison between clades that have lost spiral symmetry, like bryozoans and brachiopods, and typical spiral-cleaving clades such as annelids and molluscs, can identify the traits that were lost, or are still shared, among these groups. This comparative approach can reveal novel insights about the evolution of spiral cleavage itself, and give rise to a broader perspective of the evolutionary mechanisms underlying spiralian development.

This text is a section of my PhD thesis. Read the next section: Segmentation, a question of boundaries.


Bonner, J.T. & Bell, W.J., Jr., 1984. “What Is Money for?”: An Interview with Edwin Grant Conklin, 1952. Proceedings of the American Philosophical Society, 128(1), pp.79–84. Available at:

Child, C.M., 1900. The early development of Arenicola and Sternaspis. Wilhelm Roux’ Archiv fur Entwicklungsmechanik der Organismen, 9(4), pp.587–723. Available at:

Conklin, E.G., 1897. The embryology of Crepidula, A contribution to the cell lineage and early development of some marine gasteropods. Journal of morphology, 13(1), pp.1–226. Available at:

Costello, D.P. & Henley, C., 1976. Spiralian Development: A Perspective. American zoologist, 16(3), pp.277–291. Available at:

Dunn, C.W. et al., 2014. Animal Phylogeny and Its Evolutionary Implications. Annual review of ecology, evolution, and systematics, 45(1), pp.371–395. Available at:

Guralnick, R., 2002. A Recapitulation of the Rise and Fall of the Cell Lineage Research Program: The Evolutionary-Developmental Relationship of Cleavage to Homology, Body Plans and Life History. Journal of the history of biology, 35(3), pp.537–567. Available at:

Heath, H., 1899. The development of Ischnochiton. Zoologische Jahrbücher. Abteilung für Anatomie und Ontogenie der Tiere Abteilung für Anatomie und Ontogenie der Tiere., 12, pp.567–656. Available at:

Hejnol, A., 2010. A twist in time—the evolution of spiral cleavage in the light of animal phylogeny. Integrative and comparative biology, 50(5), pp.695–706. Available at:

Henry, J. & Martindale, M.Q., 1999. Conservation and innovation in spiralian development. Hydrobiologia, pp.255–265. Available at:

Lambert, J.D., 2008. Mesoderm in spiralians: the organizer and the 4d cell. Journal of experimental zoology. Part B, Molecular and developmental evolution, 310(1), pp.15–23. Available at:

Lambert, J.D., 2010. Developmental patterns in spiralian embryos. Current biology: CB, 20(2), pp.R72–7. Available at:

Laumer, C.E. et al., 2015. Spiralian phylogeny informs the evolution of microscopic lineages. Current biology: CB, 25(15), pp.2000–2006. Available at:

Lillie, F.R., 1895. The embryology of the Unionidae. A study in cell-lineage. Journal of morphology, 10(1), pp.1–100. Available at:

Maienschein, J., 1978. Cell lineage, ancestral reminiscence, and the biogenetic law. Journal of the history of biology, 11(1), pp.129–158. Available at:

Marcus, E., 1939. Bryozoarios Marinhos Brasileiros III. Boletim da Faculdade de Filosofia, Ciências e Letras da Universidade de São Paulo, Zoologia, 3, pp.113–299.

Mead, A.D., 1897. The early development of marine annelids. Journal of morphology, 13(2), pp.227–326. Available at:

Merkel, J. et al., 2012. Spiral cleavage and early embryology of a loxosomatid entoproct and the usefulness of spiralian apical cross patterns for phylogenetic inferences. BMC developmental biology, 12(1), p.11. Available at:

Pennerstorfer, M. & Scholtz, G., 2012. Early cleavage in Phoronis muelleri (Phoronida) displays spiral features. Evolution & development, 14(6), pp.484–500. Available at:

Riedl, R.J., 1969. Gnathostomulida from America. Science, 163(3866), pp.445–452. Available at:

Schleip, W., 1929. Die Determination der Primitiventwicklung, ein zusammenfassende Darstellung der Ergebnisse über das Determinationsgeschehen in den ersten Entwicklungsstadien der Tiere, Leipzig: Akademische Verlagsgesellschaft m.b.h. Available at:

Struck, T.H. et al., 2014. Platyzoan paraphyly based on phylogenomic data supports a noncoelomate ancestry of Spiralia. Molecular biology and evolution, 31(7), pp.1833–1849. Available at:

Wilson, E.B., 1892. The cell-lineage of Nereis. A contribution to the cytogeny of the annelid body. Journal of morphology, 6(3), pp.361–466. Available at:

  1. Also nicknamed cellular bookkepping, as recalled by E.G. Conklin: “…I followed individual cells through the development, followed them until many people laughed about it; called it cellular bookkeeping.” (Bonner and Bell, 1984, p. 81).
  2. “What a wonderful parallel is this between animals so unlike in their end stages! How can such resemblances be explained?” (Conklin, 1897, p. 195).
  3. “…if these minute and long-continued resemblances are of no systematic worth, and are merely the result of extrinsic causes, as is implied, then there are no resemblances between either embryos or adults that may not be so explained.” (Conklin, 1897, p. 195).
articles biology

Larvae as the epitome of evolution

Read the previous section: What a larva is.

Francis M. Balfour set the pace on discussions about the evolutionary importance of larvae by addressing many of the fundamental questions regarding larval evolution (Balfour, 1874; Balfour, 1880; Balfour, 1881). He wondered about the ancestry of larvae. Can larvae reveal the ancestral forms of metazoans? He indicated tests to the predictions of recapitulation. Can we find a larva that corresponds to the adult of a related group? He asked whether larvae changed during evolution. How often do larval organs evolve? And what might be the underlying mechanisms for the evolution of development. What guides the maintenance or atrophy of larval organs in adult stages? (Hall and Wake, 1999).

Perhaps, the greatest conceptual advance initiated by Balfour is that larvae are subject to variation and natural selection in the same manner as the adult stage (Balfour, 1874; Balfour, 1881). In other words, he articulated the realization that evolution can occur at any developmental stage. However, if not all embryonic features represent ancestors (or ancestral traits), the foundation of the recapitulation theory is compromised. The evolutionary debate caused by larvae influenced a more informed way to make extrapolations from ontogeny to phylogeny (Hall, 2000; Hall and Wake, 1999). It was no coincidence that one of the most vehement opponents of Haeckel’s recapitulation theory was a larvae affectionate, the biologist Walter Garstang who boldly concluded that “ontogeny does not recapitulate phylogeny, it creates it” (Garstang, 1922).

Larvae of different marine invertebrates.
Larvae of a brachiopod (left), a nemertean (center) and a bryozoan (right).

Present-day research shows that larval traits are evolutionary labile, and often correlate to ecological, developmental and other life-history factors (Strathmann and Eernisse, 1994). Evidence from diverse taxa, including gastropods (Collin, 2004), sea urchins (Raff and Byrne, 2006), ascidians (Jeffery and Swalla, 1992), sea stars (Byrne, 2006; Hart et al., 1997), nemerteans (Maslakova and Hiebert, 2014) and polyclad flatworms (Rawlinson, 2014), indicates that larval forms were modified, gained or lost in different lineages independently, and that the observed similarities are likely the result of convergent evolution.

These observations undermine scenarios about animal evolution that require the homology of larval characters (Jägersten, 1972; Nielsen, 1998; Nielsen, 2001; Nielsen, 2009; Peterson and Cameron, 1997) and are more consonant with the multiple independent evolution of metazoan larvae from a direct-developing ancestor (Page, 2009; Raff, 2008; Sly et al., 2003; Wray, 1995). Yet, the homology of larval characters such as the apical organ (e.g., Hunnekuhl and Akam, 2014; Marlow et al., 2014) or ciliated bands (e.g., Henry et al., 2007; Rouse, 1999) continues to be a central and lively discussed topic. For all the reasons above, larvae are a scandalous epitome of evolution, and the diversity of larval body patterns in marine invertebrates continue to provide a rich framework for evolutionary studies.

This text is a section of my PhD thesis. Read the next section: Spiral cleavage, an oblique matter.


Balfour, F.M., 1874. Memoirs: A Preliminary Account of the Development of the Elasmobranch Fishes. The Quarterly journal of microscopical science. Available at:

Balfour, F.M., 1880. A Treatise on Comparative Embryology, Macmillan and Company.

Balfour, F.M., 1881. A Treatise on Comparative Embryology, Macmillan and Company. Available at:

Byrne, M., 2006. Life history diversity and evolution in the Asterinidae. Integrative and comparative biology, 46(3), pp.243–254. Available at:

Collin, R., 2004. Phylogenetic effects, the loss of complex characters, and the evolution of development in calyptraeid gastropods. Evolution; international journal of organic evolution, 58(7), pp.1488–1502. Available at:

Garstang, W., 1922. The Theory of Recapitulation: A Critical Re-statement of the Biogenetic Law. Journal of the Linnean Society of London, Zoology, 35(232), pp.81–101. Available at:

Hall, B.K., 2000. Balfour, Garstang and de Beer: The First Century of Evolutionary Embryology. American zoologist, 40(5), pp.718–728. Available at:[0718:BGADBT]2.0.CO;2.

Hall, B.K. & Wake, M.H., 1999. Chapter 1 – Introduction: Larval Development, Evolution, and Ecology. In B. K. H. H. Wake, ed. The Origin and Evolution of Larval Forms. San Diego: Academic Press, pp. 1–19. Available at:

Hart, M.W., Byrne, M. & Smith, M.J., 1997. Molecular Phylogenetic Analysis of Life-History Evolution in Asterinid Starfish. Evolution; international journal of organic evolution, 51(6), pp.1848–1861. Available at:

Henry, J.Q. et al., 2007. Homology of ciliary bands in Spiralian Trochophores. Integrative and comparative biology, 47(6), pp.865–871. Available at:

Hunnekuhl, V.S. & Akam, M., 2014. An anterior medial cell population with an apical-organ-like transcriptional profile that pioneers the central nervous system in the centipede Strigamia maritima. Developmental biology, 396(1), pp.136–149. Available at:

Jeffery, W.R. & Swalla, B.J., 1992. Evolution of alternate modes of development in ascidians. BioEssays: news and reviews in molecular, cellular and developmental biology, 14(4), pp.219–226. Available at:

Jägersten, G., 1972. Evolution of the Metazoan Life Cycle First Printing edition., Academic Press Inc.

Marlow, H. et al., 2014. Larval body patterning and apical organs are conserved in animal evolution. BMC biology, 12(1), p.7. Available at:

Maslakova, S.A. & Hiebert, T.C., 2014. From trochophore to pilidium and back again – a larva’s journey. The International journal of developmental biology, 58(6-8), pp.585–591. Available at:

Nielsen, C., 1998. Origin and evolution of animal life cycles. Biological reviews of the Cambridge Philosophical Society, 73(02), pp.125–155. Available at:

Nielsen, C., 2001. Phylum Ectoprocta. In Animal Evolution: Interrelationships of the Living Phyla. Oxford University Press, pp. 244–263.

Nielsen, C., 2009. How did indirect development with planktotrophic larvae evolve? The Biological bulletin, 216(3), pp.203–215. Available at:

Page, L.R., 2009. Molluscan larvae: Pelagic juveniles or slowly metamorphosing larvae? The Biological bulletin, 216(3), pp.216–225. Available at:

Peterson, K.J. & Cameron, R.A., 1997. Set-aside cells in maximal indirect development: Evolutionary and developmental significance. BioEssays: news and reviews in molecular, cellular and developmental biology, 19(7), pp.623–631. Available at:

Raff, R.A., 2008. Origins of the other metazoan body plans: the evolution of larval forms. Philosophical transactions of the Royal Society of London. Series B, Biological sciences, 363(1496), pp.1473–1479. Available at:

Raff, R.A. & Byrne, M., 2006. The active evolutionary lives of echinoderm larvae. Heredity, 97(3), pp.244–252. Available at:

Rawlinson, K.A., 2014. The diversity, development and evolution of polyclad flatworm larvae. EvoDevo, 5(1), p.9. Available at:

Rouse, G.W., 1999. Trochophore concepts: ciliary bands and the evolution of larvae in spiralian Metazoa. Biological journal of the Linnean Society. Linnean Society of London, 66(4), pp.411–464. Available at:

Sly, B.J., Snoke, M.S. & Raff, R.A., 2003. Who came first–larvae or adults? origins of bilaterian metazoan larvae. The International journal of developmental biology, 47(7-8), pp.623–632. Available at:

Strathmann, R.R. & Eernisse, D.J., 1994. What Molecular Phylogenies Tell Us about the Evolution of Larval Forms. Integrative and comparative biology, 34(4), pp.502–512. Available at:

Wray, G.A., 1995. Punctuated evolution of embryos. Science, 267(5201), pp.1115–1116. Available at:

biology articles

Endless larval forms most beautiful: what a larva is

This text is a section of my PhD thesis.

The Latin word lārva means evil spirit, ghost or mask1. In the 18th century, the naturalist Carolus Linnaeus was the first to employ the word larva to describe a stage in the life of an animal in which its adult form is still hidden or masked (Linnaeus, 1767, p. 534). An exemplar case of this new biological meaning is the maggot—the larval stage of a fly—whose wormy form and life style truly differs from its flying adult stage.

Not all larvae, however, are masked forms. The larval body of some marine snails2, for example, is very similar to its adult body, except for the dazzling presence of a ciliated velum, used by the larva to swim and gather food (Collier, 1997). In more general terms, larval stages are considered to be a modification of embryonic development usually characterized by a morphology and habitat that are disparate from the adult stage (Hall and Wake, 1999). Because embryonic development can change in a multitude of ways, as evidenced by the great diversity of larval forms in nature (see below), there is no precise definition of larva (Hickman, 1999; Strathmann, 1993). Thus in practice, what a larva is, is defined case by case according to the organism and to one’s research background.

The majority of animals on this planet have a complex life cycle with one or more larval stages. Collectively, marine invertebrates represent a great part of the observed larval diversity. Molluscs have the veliger, a shelled larva with the ciliated velum mentioned above; echinoderms have the pluteus, a spaceship-like larva with eight food-capturing arms, and the brachiolaria, a free-swimming larva driven by body-length dancing arms; bryozoans have the cyphonautes, a paper-thin triangular larva that sails over kelp blades; crustaceans have the zoea, an armored larva that swims as if using a jet pack; nemerteans have the pilidium, a larva with lobes and lappets in the form of a deerstalker cap… and this list goes on. The diversity of larval forms is astonishing.

Sample of the diversity of metazoan larval forms. Larvae are not to scale. Photos from the Cifonauta marine biology image database (Migotto and Vellutini, 2011).
Sample of the diversity of metazoan larval forms. Larvae are not to scale. Photos from the Cifonauta marine biology image database (Migotto and Vellutini, 2011).

Most of these charismatic larval figures were discovered in the 19th century by the naturalist founders of comparative embryology (Hall and Wake, 1999). At the time, the ideas of Karl Ernst von Baer and Ernst Haeckel had great influence on the understanding of embryonic development (Guralnick, 2002; Hall, 2000). Ontogeny was seen as the unfolding of an immutable process that represents the evolutionary history of an organism—an idea known as recapitulation or Haeckel’s biogenetic law: “ontogeny is a rapid and shortened recapitulation of phylogeny.” (Gould, 1977; Haeckel, 1866).

These influential ideas were directly challenged by the mere existence of larvae. Or more generally, challenged by the existence of differentiated developmental stages that are, at the same time, functionally adapted to their environment and morphologically diverse. Such impressive variety of larval forms instigated questions about the relationship between the embryonic development of an individual (ontogeny) and the evolutionary history of a lineage (phylogeny).

Do larvae represent ancestral adult forms? How many times have larvae evolved? Are larval structures homologous or independently evolved? Soon, there was an urge to rationalize the diversity of larval forms into an evolutionary context.

Read the next section: Larvae as the epitome of evolution.


Collier, J.R., 1997. Gastropods, the Snails. In S. F. Gilbert & A. M. Raunio, eds. Embryology: constructing the organism. Sinauer Associates, Inc., pp. 189–217.

Gould, S.J., 1977. Ontogeny and phylogeny, Harvard University Press.

Guralnick, R., 2002. A Recapitulation of the Rise and Fall of the Cell Lineage Research Program: The Evolutionary-Developmental Relationship of Cleavage to Homology, Body Plans and Life History. Journal of the history of biology, 35(3), pp.537–567. Available at:

Haeckel, E., 1866. Generelle Morphologie der Organismen, Georg Reimer, Berlin. Available at:

Hall, B.K., 2000. Balfour, Garstang and de Beer: The First Century of Evolutionary Embryology. American zoologist, 40(5), pp.718–728. Available at:[0718:BGADBT]2.0.CO;2.

Hall, B.K. & Wake, M.H., 1999. Chapter 1 – Introduction: Larval Development, Evolution, and Ecology. In B. K. H. H. Wake, ed. The Origin and Evolution of Larval Forms. San Diego: Academic Press, pp. 1–19. Available at:

Hickman, C.S., 1999. Chapter 2 – Larvae in Invertebrate Development and Evolution. In B. K. H. H. Wake, ed. The Origin and Evolution of Larval Forms. San Diego: Academic Press, pp. 21–59. Available at:

Linnaeus, C., 1767. Systema Naturæ, Impensis direct. Laurentii Salvii. Available at:

Migotto, A.E. & Vellutini, B.C., 2011. Cifonauta – marine biology image database. Cifonauta, an image database for marine biology. Available at: [Accessed December 16, 2015].

Sars, M., 1837. Beitrag zur Entwicklungsgeschichte der Mollusken und Zoophyten. Archiv für Naturgeschichte, 3, pp.402–407. Available at:

Strathmann, R.R., 1993. Hypotheses on the Origins of Marine Larvae. Annual review of ecology and systematics, 24, pp.89–117. Available at:

Young, C.M., 1990. Larval ecology of marine invertebrates: A sesquicentennial history. Ophelia, 32(1-2), pp.1–48. Available at:

  1. American Heritage® Dictionary of the English Language, Fifth Edition. (2011). Accessed November 13 2015 at
  2. Michael Sars, one of the Norwegian biologists giving the name to the Sars Centre, was among the first to describe the development of molluscs from a swimming larva (Sars, 1837; Young, 1990).
science articles code

Search PLOS articles using DuckDuckGo

Last year I decided to experiment with DuckDuckHack, the developer platform for the search engine DuckDuckGo. The idea was to use the instant answers to find scientific articles as a quick Google Scholar shortcut.

It’s feasible, in principle, but I decided to try something simpler. A plugin that uses the PLOS API to search their articles and display in the instant answer box.

To use it you just need to add the word “plos” + keywords (example above). The result is a list of titles and dates of the five most-relevant articles with direct links. Hovering the mouse over a link will show the authors and which PLOS journal. This final format was simplified after the initial pull request and polished up in the second.


The code is a simple Perl function that connects the PLOS API to DuckDuckGo, and a javascript function that handles the search response. The DDG community was quite friendly to help out with the code.

Since DuckDuckGo is less used than Google I guess the number of users might be low. Maybe I’m the only one… It would be amazing if it could query the whole scientific literature! But well, I like this little hack. I guess it’s the excitement of connecting services using APIs.