How to achieve cellular replication without fail: lessons from bacterial cells.

Speaker: Christine Jacobs-Wagner
Department: Microbial Sciences Institute, Yale West Campus
Subject: How to achieve cellular replication without fail: lessons from bacterial cells.
Location: TU Delft (BN Seminar)
Date: 13-10-2017     

 Author: Nemo Andrea

 The topic of today’s talk was cellular replication, which, in Christine’s opinion, is the ability that separates the living from the non-living. In order to study this process, they study bacterial replication, as bacteria both divide rapidly and do so with high accuracy. The speaker stressed how, if one stops to think about it, achieving such short (20 minutes) division times in a varying environment with in the inherently highly stochastic environment of a living cell, is a truly remarkable feat. Thus, the robustness of bacterial replication and the relative simplicity of prokaryotic systems as compared to eukaryotic model systems make bacteria the candidate of choice for her research.

As there are many topics that can be explored within cellular replication, the speaker decided to focus on a single system that her group had recently done work on. Many bacteria have plasmids, which can contain vital functions for a bacterium, thus requiring efficient segregation of plasmids upon division. While plasmids diffuse throughout the cell and are thus, on average, equally divided among the two halves of a bacterium, suggesting that in the case of cell division no problem should arise, this is not the case. If a plasmid is present in high copy number, the chance of one daughter cell having significantly fewer copies of the plasmid is very small, but if a plasmid has low copy number the probability of one daughter cell ending up with no copies due to low number noise becomes significant. It is thus that bacteria have developed methods to effectively separate the plasmids that are present in low numbers. One could argue that it is not the bacterium driving the evolution of such a system but rather the plasmid itself, as plasmids that effectively do this ensure their survival, but that is really more a matter of perspective than anything else, and beside the point argued in this talk.

The plasmid that the group focused on displayed very curious behavior. The plasmids (in elongated bacteria) are distributed equidistantly along the long axis. The plasmids do diffuse, but stay in roughly the same area over time. It should be apparent that if such a distribution is maintained, the plasmids will be divided equally among daughter cells. As it turns out, this is achieved through a particular variant of the Par system. This variant uses just two proteins: PAR A/B, which are encoded on the plasmid itself. ParB binds to the plasmid, and does this by recognizing specific sequences on the plasmid, ensuring selective binding. ParA, on the other hand, binds to ParB, after which ParB will stimulate the ATPase activity of ParA, which will then unbind from the DNA. ParA also unspecifically binds to the DNA (of the bacterium). They observed (in case of a single plasmid) an oscillation of ParA from one side of the cell to the other, with the plasmid (with ParB) following the ParA signal. They then produced a first model, to test if simple Brownian dynamics of a diffusing plasmid could reproduce such behavior. They found that this was insufficient and concluded from this that a translocating force must be present to create this behavior.

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Figure taken from: DNA-relay mechanism is sufficient to explain ParA-dependent intracellular transport and patterning of single and multiple cargos. ; cropped

The key insight that was missing from the first simulation was the fact that DNA is not static, but moves around in the cell. They determined that the movement of a locus in the DNA can essentially be seen as an elastic harmonic potential, with a force of around 0.04pN. This is a really small force, even by cellular standards. They redid the simulations with this idea incorporated and found that they were able to reproduce the behavior observed in bacteria. The conceptual change is now as follows: The plasmid with ParB is pulled around through its multiple connections with multiple harmonic potentials through ParA. It will then unbind from the ParA in that area and move in a direction (slightly). In the new situation, the plasmid will ‘see’ more ParA in the direction of movement than behind it (which is the previous location, where it made ParA unbind) essentially creating a gradient of forces in the direction of movement. The speed of the plasmid seemed to correspond well with the force of the harmonic potentials. The initial symmetry breaking is caused by the stochastic nature of ATP hydrolysis.

The speaker mentioned another recent model that was developed that may explain similar phenomena, but pointed out that certain parts of this specific system are most likely not reconcilable with that model. The model presented today was also extremely elegant requiring just two components; one that binds to chromosomes, and another that modulates that first component’s lifetime. I quite enjoyed the talk, as Christine was an excellent presenter. While I am generally not easily convinced by simulations, this model seems to make sense both mechanistically and matches real world experiments. We also had a brief session afterwards to ask some questions, which was cut short due to unforeseen circumstances. I do appreciate her taking the time to sit down with us and answer some questions – even the ones not related to the talk.


Targeting Epigenetic Changes in Immune Cells: Implications in Disease

Speaker: Esteban Ballestar
Department: Bellvitge Medical Research Institute (IDIBELL), Barcelona Spain
Location: Erasmus MC
Date: July 13, 2017
Author: Teun Huijben

Esteban Ballestar got his Bachelor and Master degree at the University of Valencia in Spain, followed by a PhD. Afterwards he did a Postdoc abroad and returned to start his own research group at the same university. The main interest of his group is the DNA methylation, and especially in the context of diseases involving the immune system.

The first part of Estebans talk was about mapping DNA methylation in immunological diseases to understand which proteins are involved in the disease. The group of diseases they studied were Common Variable Immunodeficiencies Diseases (CVID) in which the body has not enough primary antibodies. These diseases are mostly caused by severe deficiencies in the number of switched memory B-cells. With switched B-cells we mean activated B-cells that start producing the antibodies in high quantities after recognizing the antigen. By mapping the DNA methylations of these B-cells, they hope to find genes that are differently methylated and are mostly likely causing the disease.

Methylation of DNA means that a methyl group is added to the 5-prime end of a cytosine (5mC) nucleobase. This can only be done if the cytosine is next to a guanine (see Figure 1). DNA methylation is maintained by de-novo DNA methyltransferases (mostly DNMT1, DNMT3A and DNMT3B). DNA methylations can be actively removed by demethylation, in which the 5mC is oxidized to a 5hmC or 5caC. Adding or removing methyl group to the DNA has an effect on the gene expression of that particular gene.


Figure 1: DNA methylation. The cytosine of the CG-pair gets methylated by a de-novo methyltransferase (DNMT). [S. Zakhari 2015]

To identify disease causing genes, Estebans group did DNA methylation profiling of the B-cells from a CVID patient. To eliminate as much side effects as possible, thy only investigated twins of which one sibling had CVID and the other was healthy. After collecting the B-cells, they performed DNA methylation profiling and looked at genes with different methylation profiles between the two brothers. They found 230 genes that are more methylated in the CVID patient and 81 genes that are less methylated. Gene ontology analysis showed that most of the genes were related to immune responses, indicating that changing the gene expression of these genes can cause an immune related disease.

All the genes that showed different methylation profiles between healthy and CVID patient, were taken into further research. The B-cells are sorted on the fact whether they were naive (not yet switched to active) or switched. They found that in healthy persons most genes got demethylated after the transformation from naive to switched. On the other hands, the same genes in CVID patients showed no decrease in methylation, a second indication that these genes are involved in the disease.

However, the question remains whether the different methylation profile itself causes the disease, or is it a downstream effect caused by other factors. To investigate this, more research needs to be done on this subject. Also, more methylation profiles of twins are needed to draw real conclusions about the disease causing genes, since one set of results of the statistically valid enough. Overall, the talk of Esteban was interesting and he is a very good speaker. Despite using many difficult immunology term, he explained very clear the research his lab is doing.

Disrupting ultrashort nucleic acid duplex with mechanical force

Speaker: Kevin Whitley

Department: University of Illionois at Urbana -Champain

Subject: Disrupting ultrashort nucleic acid duplex with mechanical force

Location: Bionanoscience Department, TU Delft

Date: 30-06-2017

Author: Mirte Golverdingen

Kevin Whitley did research to the hybridization kinetics of DNA under force. He tried to understand the transition state of nucleic acid hybridization. Hybridization is used for gene manipulation and targeting for DNA nanomachines, tweezers and bipadal walkers. Short strands (about 10 bp) show a ‘all of nothing’ behavior when hybridizing. Fist one or two base pairs bind and then all base pairs bind without stable intermediates. This describes a two-state reaction in a schematic energy landscape. The kinetics of any of these states is determined by the transition state between them. Whitley searched for the transition state between the bound and the unbound DNA.

To research the hybridization kinetics of the DNA the end-to-end extension of the DNA was researched. This is the length of the top of the energy to the bottom of the energy where the DNA is hybridized. Bell’s equation describes the energy barrier between the end-to-end extension. So, by measuring the rates of the Bell’s function you can calculate the end-to-end distance.

Whitley measured the hybridization rates by using optical beads that are bound to the DNA by double stranded handles and with a 9 nucleotides long middle strand. This short part is the binding site for short DNA. The distance between the two beads was approximately one. By using a flow chamber Whitley was able to look at oligo that were free in solution. He was able to measure the koff and kon rate under a constant amount of force. In this way, he could measure the moment a ss DNA oligomer binds to the middle strand by measuring the position of the bead. The ds DNA is stiffer than ss DNA, and therefore the position of the bead changes. By using this method, Whitley could measure the difference between ss and ds DNA up until a force of 12 pN.

By using Fleezers, which is a combination between fluorescence and optical tweezers, Whitley was able to label the oligomer with a fluorophore (see figure 1). In this way, also forces lower than 12 pN can be measured. Again, they calculated the binding and unbinding rates of the DNA. While the unbinding rate of the DNA goes up linearly when the force increases, the binding rate also goes up, however, not linearly. When the oligomer becomes longer, the unbinding rate goes down. For the binding rate no dependence on the length of the oligomer can be found.


Figure 1: Measurement of single-oligonucleotide hybridization kinetics under force. (A) Schematic of the hybridization assay (not to scale). An engineered DNA molecule (red) containing a short, central ssDNA region flanked by long double-stranded DNA (dsDNA) handles is held under constant force by polystyrene beads (grey spheres) held in optical traps (orange cones). A fluorescence excitation laser (green cone) is focused on the central ssDNA region. Short oligonucleotides (blue) labeled with a Cy3 fluorophore at the 3΄ end (green disk) bind and unbind to the complementary ssDNA sequence in the center of the tethered DNA. The binding and unbinding is observed by the fluorescence emitted from the attached fluorophores.

Adapted from: Kevin D. Whitley, Matthew J. Comstock, Yann R. Chemla; Elasticity of the transition state for oligonucleotide hybridization. Nucleic Acids Res 2017; 45 (2): 547-555. doi: 10.1093/nar/gkw1173

When looking at Bell’s equation, the binding rate should be an exponential force-dependence, however, non-exponential behavior is reproducible across other conditions. The end-to-end distance is therefore not constant, and the Bells’ equation changes so an integral is used instead.  By calculating the unbound and bound extension by using the worm like chain model for double stranded DNA they were able to calculate the end-to-end extension. In this way, they also obtained the persistence length and contour length of the end-to-end extensions. He repeated these calculation for all different lengths of oligomers. The transitions state persistence length is comparable to ssDNA, however it is a bit stiffer.

The transition state is probably no random process, as simple polymer estimations predict a low probability of spontaneous alignments. The single strand that is a bit stiffer, as measured in the transition state, this could correspond to prearranging of the strands, prior to nucleation. Some enzymes can pre-organize mRNA so the transition state energy barrier becomes lower. Moreover, fast target finding is possible when there are preorganized bases by enzymes. In this way, they can speed hybridization up by pre-paying the entropy penalty.

Whitley told a clear story on obtaining the end-to-end distance of the DNA hybridization translation state. The combination of Optical Tweezers and fluorescence was very cool. Moreover, using the mathematics in combination with the biology inspired me as Nanobiology student.

Evolution and Assembly of Eukaryotic Chromatin

Speaker: Fransesca Mattiroli

Department: Lugi Lab, University of Colorado Boulder

Subject: Evolution and assembly of eukaryotic Chromatin

Location: TU Delft, Bionanoscience department

Date: 10-02-2017

Author: Mirte Golverdingen


Fransesca Mattiroli’s research is focussed on the DNA packaging units called nucleosomes. These structures organize DNA in the eukaryotic cell nucleus. Nucleosomes are formed by an octameric complex of folded histone dimers called the H3-H4 and H2A-H2B dimers. In mammals, the histones have histone tails which highly contribute to post-translational modifications and they stabilize the nucleosome. Nucleosomes need to assemble and disassemble when they bind to the genome DNA. Histone modifications and variants are dynamic and can promote or inhibit certain interactions. The nucleosome dynamics and compositions have a direct effect on transcription, translation and repair.

The first main interest of Mattiroli is the evolutionary origin of the nucleosome. The nucleosomes are very well conserved through species. Mattiroli focusses on the structural conservation of the histone dimers in Archaea. They, however, miss the tails that contribute to post-translational modification. So, how do these species organize their archaeal genome?

The archaeal histone binding to DNA is similar to eukaryotic histone binding. Archael histones, however, do not form octamers. They can form a much longer structure instead, called nucleosomal ramps. In Vivo, this structure also forms, the longest ramp they found was 90 bp long. So, they found a new way of arranging histone DNA complexes.

Histones are formed on the DNA in two steps, first, two H3-H4 dimers form a tetrasome, then two H2A-H2B dimers attach to this tetrasome forming a nucleosome. Histone chaperones shield the charges of the histones and facilitate their deposition on DNA. However, not much is known on how the chaperones actually contribute to this deposition step. The Chromatin Assembly Factor 1, CAF-1, is Mattiroli’s main interest. CAF-1 mediates in this histone deposition step and is essential in multicellular organisms. Matteroli tried to understand how CAF-1 contributes to the deposition step.

Mattiroli’s first step was to research how CAF-1 binds the H3-H4 dimer. She used mass spectrometry (HX-MS) with a hydrogen-deuterium exchange. She could, in this way, measure the change in mass and which regions have the largest changes in deuterium uptake. This region could then be the binding site of CAF-1 on H3-H4. When CAF-1 binds to the dimer, they see a stabilization of the dimer. This result indicates the following hypothesis: Only if a H3-H4 dimer is bound to CAF-1 it can form a tetrasome.

A next step for Mattiroli was to test if CAF-1 can form nucleosomes in vitro, in absence of other proteins. To test this, Mattiroli mixed CAF-1, histones and DNA, treat them with micrcococcal nuclease to digest unprotected DNA and purified and quantified the length of DNA covered by histones. The result showed that CAF-1 is able to assemble tetrasomes, and therefore enabling nucleosome formation in vitro.

So, how is the H3-H4 tetrasome on the DNA formed? Mattiroli used increased lengths of DNA, to trap any intermediates in the process. Mattiroli showed that the forming of the H3-H4 dimer activates the DNA binding of the dimer. The key intermediate that mediates the DNA binding results to be two CAF-1 units. This was the most interesting result so far, because it was never showed before that two independent CAF-1 were involved in the H3-H4 DNA binding.

The interesting and clear seminar showed again how complex the system of DNA and all the DNA-interacting molecules is. The research of Mattiroli gives a good foundation for more research to nucleosomes and their interaction with DNA. Bringing us closer to fully understand the biological system of DNA.

Honours 7

Figure 1. Canonical and variant nucleosomes

(A) Elements of the histone fold and structures of Xenopus leavis H2A–H2B, H3–H4 and (H3–H4)2 (PDB ID: 1KX5). (B) Structure of the canonical Xenopus leavis nucleosome (PDB ID: 1KX5). Other nucleosome structures, such as the human nucleosome, are structurally similar. (C) Structure of the CenH3CENP‐A‐containing nucleosome (PDB ID: 3AN2). (D) Zoomed view of the αN helix of CenH3CENP‐A (left) and H3 (right) involved in stabilizing the DNA ends. Histone H3 is blue, CenH3CENP‐A is cyan, H4 is green, H2A is yellow, H2B is red, and DNA is white.

Adapted From: Mattiroli, F., D’Arcy, S., & Luger, K. (2015). The right place at the right time: chaperoning core histone variants. EMBO reports, e201540840.

Nanotechnology for biophysics: from single molecules towards synthetic cells

Speaker: Cees Dekker
Nanotechnology for biophysics: from single molecules towards synthetic cells
Location: Lecture room E (TU Delft)
Date: 09-02-2017

Kristian Blom

After six months of following theoretical physics and astronomy courses for my minor, I was looking forward to hear something about nanobiology again. Therefore I took the opportunity to visit a seminar given by Cees Dekker for the quantum nanoscience department about nanotech nology for biophysics.

To break the ice between the applied physicists (audience) and the nanobiologist,  prof. Dekker started his talk with a quick review of his scientific career. In 1984 he started his career in solid state physics (which impressed the audience for sure) and over the years he got more and more fascinated by biophysics and nanobiology. Therefore he made the decision to switch from quantum physics to biophysics around 2000. In the past 15 years Prof. Dekker mainly focused on the study of cellular components using the top-down approach. For this kind of research nanotechniques are used to probe the biological cell at its most fundamental level; single biomolecules.

So what is nanotechnology? Specifically in the field of biophysics we can describe nanotechnology as a toolbox of instruments for studies at the single-atom and single-molecule level. A very famous example of this is the scanning tunneling microscope. Another example are the optical and magnetic tweezer with which you can measure the mechanical properties of DNA. Prof. Dekker explained the audience that with a magnetic tweezer you can coil up a DNA strand by applying an external rotating magnetic field to a bead with one end of a DNA molecule is attached. 

Currently the research group of prof. Dekker focuses on three main topics: Single-molecule biophysics of DNA and DNA-protein complexes, solid-state nanopores and its applications, and finally bacteria in nanostructures. I will discuss the former topic in a bit more detail by explaining one of the recent findings of  the Dekker group in this field.

Figure 1 – Intercalation-induced Supercoiling of DNA (ISD) Source: Ganji, M.; Hyun Kim, S.; van der Torre, J.; Abbondanzieri, E.; Dekker, C. Nano Lett, 2016, 16 (7), pp 4699-4707

The subject of this recent finding is on DNA supercoiling, which is the over- (positively coiled) or under-winding (negatively coiled) of the DNA strand. Since supercoiling plays a very important role in biological processes such as DNA compaction, gene expression and DNA replication, it is very important for the cell to keep control of the supercoiling state of DNA. A specific kind of supercoiled DNA configuration is the plectoneme; a DNA helix which is coiled onto itself. By using an intercalating dye the Dekker group induced supercoils within a linear DNA molecule that was bound to a surface at its two ends. Thereafter they investigated the plectoneme position and dynamics using epifluorescence microscopy. What they found is that the observed plectoneme density and the nucleation and termination rates showed position-dependent variations. More specifically, they found a strong peak in the plectoneme density, nucleation and termination rate near one end of the DNA (see figure 2). As nucleation of a plectoneme requires a lot of bending energy, a locally region where the DNA is already bend a little bit (intrinsic curvature) by external structures (e.g. DNA-bound proteins) would significantly promote the formation and growth of a plectoneme at this site. Therefore the Dekker group inserted a 10-nucleotide mismatched sequence in the middle of the DNA strand. They observed that the plectoneme density showed a peak at the position of the mismatched sequence. So, as expected, flexible DNA bubbles promote the plectoneme formation.

Figure 2 – DNA sequence-dependent pinning of plectonemes. (A) Plectoneme densities of 46 identical DNA molecules (thin lines) and their average (red thick line). (B) Averaged nucleation (orange) and termination (blue) rates observed. C) Averaged position-dependent plectoneme size distributions. Source: Ganji, M.; Hyun Kim, S.; van der Torre, J.; Abbondanzieri, E.; Dekker, C. Nano Lett, 2016, 16 (7), pp 4699-4707

My heart leapt for joy when prof. Dekker mentioned that a theoretical model will be made about the nucleation and termination of plectonemes. Although experimental data tells us more about the truth (if the experiment is done properly) than a theoretical model, I think you can only understand a physical phenomenon in the very detail when someone can explain it using the fundamental laws of nature. After this very interesting part of the talk prof. Dekker quickly went over the other two topics where is research group is interested in. At the very end someone from the audience asked if quantum mechanics plays a significant role in the functioning of a cell. The answer of prof. Dekker was that it isn’t that important for phenomenon in the cell (except for photosynthesis and energy conversion). This disappointed me a little bit, since one of my future plans is to use quantum mechanics to get a better understanding of the cell (therefore I will also follow quantum mechanics in the coming semester). But luckily not everything is discovered so far, so for me there is still hope that one day quantum mechanics will be an essential part of every cell biology course!

3D mammalian genome organization and function

Speaker: Wouter de Laat

Department: Bionanoscience

Subject: 3D mammalian genome organization and function          

Location: TU Delft

Date: 11-2-16

Author: Hielke Walinga

Our DNA consists of long strands of bases that appear to have no function at all. Only 3 % of the DNA is actually consisting genes. A long time researchers thought only the genes of the DNA mattered. However, it becomes more and more clear that the junk DNA hides a lot of information related to the expressing of these genes. Not only did they turn out to consist of a large landscape of regulatory elements, they also contain parts which guide to fold the DNA in a specific 3D construction which is also influencing the expressing pattern. Wouter de Laat investigated this 3D construction and talked about methods to reveal this, but also talked about a mechanism explaining the self-organization of this 3D construction.

When taking into account that junk DNA surrounding the genes are important to the expressing of the genes, one can deduce that the position of these genes in the DNA is important for the expressing pattern. A good way to test this is to place genes at random positions in the DNA. The Spitz lab executed these kinds of experiments and indeed showed that the expressing patterns can sometimes change very much.

A way to explain this is given by the hypothesis that when DNA strands are looping, the gene and its enhancer are placed close to each other in space. To test this hypothesis, a technique called Chromosomes Conformation Capture (referred to as 3C) is developed. This technique makes use of formaldehyde proteins which connect to the DNA, creating hairballs of DNA. These hairballs are the loops of DNA. When this is digested, the remaining DNA is all in small parts which used to be the loop. In the next step, it uses PCR with primers linking to the genes and its enhancer. The PCR then reveals if the gene and its enhancer were located in the same loop.

Other more elaborate techniques of the previous mentioned technique are the 4C and the Hi-C method. In the 4C method the topological distance between one locus is measured to all other DNA. The Hi-C method on the other hand measures all loci to all other loci. This method reveals beautifully how DNA is topologically organized.

Next, Wouter de Laat wanted to explain us about the construction of these loops. His hypothesis stated that on the root of the loop there are two CTCF anchors recruiting condensin. This condensin links the strands together creating the loop. To prover this hypothesis, researchers removed one anchor and observed that no loop is formed. (This was then tested with the 3C technique.) Another experiment showed a more surprising result. When one anchor was switched the loop was also not created.

At first, this made sense, but at the end of the presentation somebody made an important note. In a three dimensional world, switching one anchor does not automatically result in a prevention of linking. Especially when the loop is very large, this would make no difference. An important feature Wouter de Laat missed in his presentation was the coiling and super-coiling of the DNA. He missed to mention and elaborate on the physics of DNA. A hole in biology Nanobiology is hoping to fill.

Image 1: A heat map showing the topological distances in chromosome 14. This was created by the Hi-C technique. (Source: Lieberman-Aiden E, van Berkum NL, Williams L, Imakaev M, Ragoczy T, Telling A, Amit I, Lajoie BR, Sabo PJ, Dorschner MO, Sandstrom R, Bernstein B, Bender MA, Groudine M, Gnirke A, Stamatoyannopoulos J, Mirny LA, Lander ES, Dekker J,Comprehensive mapping of long-range interactions reveals folding principles of the human genome, Science 326, 289–293, 2009)



Coordination of daughter strand synthesis

Single replication machines at work:

The coordination of daughter strand synthesis

By Karl Duderstradt (van Oijen lab, University of Groningen)

One of the most important events a cell has to undergo before it will divide is DNA replication. In this event, the amount of genomic DNA is doubled by taking the two strands apart and synthesize complementary strands to them. This process is schematically shown in Figure 1:

Replication machinery

Figure 1: Figure 5-19 in Molecular Biology of the Cell (©Garland Science)

The collection of molecules involved in the replication machinery called the replisome. It is not yet entirely known how the replication machinery is assembled and what coordinates the function of it, but Karl Duderstadt works on these problems. He does this by single-molecule assays, in which single molecules are labeled with a fluorescent dye or protein, and can be followed. The power of single molecule assays is that instead of looking at what a lot of particles do together and identify averages, you can look at individual rates. This is similar to the difference between looking down from a skyscraper down to traffic and standing beside the road looking at individual cars. Every car will do very different things in both cases, but when you look down from the skyscraper you only see the average of many.

Karl uses E. Coli in his research. To be more specific, he looked at the bacteriophage T7 replisome, and was especially interested in how the enzymatic events on leading and lagging strands were coordinated. To better understand his findings, take a look at Figure 2 which shows the replication slightly more detailed. As can be seen there, the replication machinery uses two kinds of loops to synthesize the lagging-strand. These loops are called the priming loop and the trombone or replication loop. About 1% of the lagging strand DNA is synthesized in the replication loop, which are also five times less likely than the priming loops.

Trombone and priming loops

Figure 2: Priming loop and Trombone loop. Found at

Kasper Spoelstra –