Category Archives: Group Meetings

What we discuss during cake at our Tuesday afternoon group meetings

Using RDKit to load ligand SDFs into Pandas DataFrames

If you have downloaded lots of ligand SDF files from the PDB, then a good way of viewing/comparing all their properties would be to load it into a Pandas DataFrame.

RDKit has a very handy function just for this – it’s found under the PandasTool module.

I show an example below within Jupypter-notebook, in which I load in the SDF file, view the table of molecules and perform other RDKit functions to the molecules.

First import the PandasTools module:

from rdkit.Chem import PandasTools

Read in the SDF file:

SDFFile = "./Ligands_noHydrogens_noMissing_59_Instances.sdf"
BRDLigs = PandasTools.LoadSDF(SDFFile)

You can see the whole table by calling the dataframe:

BRDLigs

The ligand properties in the SDF file are stored as columns. You can view what these properties are, and in my case I have loaded 59 ligands each having up to 26 properties:

BRDLigs.info()

It is also very easy to perform other RDKit functions on the dataframe. For instance, I noticed there is no heavy atom column, so I added my own called ‘NumHeavyAtoms’:

BRDLigs['NumHeavyAtoms']=BRDLigs.apply(lambda x: x['ROMol'].GetNumHeavyAtoms(), axis=1)

Here is the column added to the table, alongside columns containing the molecules’ SMILES and RDKit molecule:

BRDLigs[['NumHeavyAtoms','SMILES','ROMol']]

Confidence (scores) in STRING

There are many techniques for inferring protein interactions (be it physical binding or functional associations), and each one has its own quirks: applicability, biases, false positives, false negatives, etc. This means that the protein interaction networks we work with don’t map perfectly to the biological processes they attempt to capture, but are instead noisy observations.

The STRING database tries to quantify this uncertainty by assigning scores to proposed protein interactions based on the nature and quality of the supporting evidence. STRING contains functional protein associations derived from in-house predictions and homology transfers, as well as taken from a number of externally maintained databases. Each of these interactions is assigned a score between zero and one, which is (meant to be) the probability that the interaction really exists given the available evidence.

Throughout my short research project with OPIG last year I worked with STRING data for Borrelia Hermsii, a relatively small network of scored interactions across 815 proteins. I was working with v.10.0., the latest available database release, but also had the chance to compare this to v.9.1 data. I expected that with data from new experiments and improved scoring methodologies available, the more recent network would be more or less a re-scored superset of the older. Even if some low-scored interactions weren’t carried across the update, I didn’t expect these to be any significant proportion of the data. Interestingly enough, this was not the case.

Out of 31 264 scored protein-protein interactions in v.9.1. there were 10 478, i.e. almost exactly a third of the whole dataset, which didn’t make it across the update to v.10.0. The lost interactions don’t seem to have very much in common either — they come from a range of data sources and don’t appear to be located within the same region of the network. The update also includes 21 192 previously unrecorded interactions.

densityComparison

Gaussian kernel density estimates for the score distribution of interactions across the entire 9.1. Borrelia Hermsii dataset (navy) and across the discarded proportion of the dataset (dark red). Proportionally more low-scored interactions have been discarded.

Repeating the comparison with baker’s yeast (Saccharomyces cerevisiae), a much more extensively studied organism, shows this isn’t a one-off case either. The yeast network is much larger (777 589 scored interactions across 6400 proteins in STRING v.9.1.), and the changes introduced by v.10.0. appear to be scaled accordingly — 237 427 yeast interactions were omitted in the update, and 399 836 new ones were added.

discardedYeast

Kernel density estimates for the score distribution for yeast in STRING v.9.1. While the overall (navy) and discarded (dark red) score distributions differ from the ones for Borrelia Hermsii above, a similar trend of omitting more low-scored edges is observed.

So what causes over 30% of the scored interactions in the database to disappear into thin air? At least in part this may have to do with thresholding and small changes to the scoring procedure. STRING truncates reported interactions to those with a score above 0.15. Estimating how many low-scored interactions have been lost from the original dataset in this way is difficult, but the wide coverage of gene co-expression data would suggest that they’re a far from negligible proportion of the scored networks. The changes to the co-expression scoring pipeline in the latest release [1], coupled with the relative abundance of co-expression data, could have easily shifted scores close to 0.15 on the other side of the threshold, and therefore might explain some of the dramatic difference.

However, this still doesn’t account for changes introduced in other channels, or for interactions which have non-overlapping types of supporting evidence recorded in the two database versions. Moreover, thresholding at 0.15 adds a layer of uncertainty to the dataset — there is no way to distinguish between interactions where there is very weak evidence (i.e. score below 0.15), pairs of proteins that can be safely assumed not to interact (i.e. a “true” score of 0), and pairs of proteins for which there is simply no data available. While very weak evidence might not be of much use when studying a small part of the network, it may have consequences on a larger scale: even if only a very small fraction of these interactions are true, they might be indicative of robustness in the network, which can’t be otherwise detected.

In conclusion, STRING is a valuable resource of protein interaction data but one ought to take the reported scores with a grain of salt if one is to take a stochastic approach to protein interaction networks. Perhaps if scoring pipelines were documented in a way that made them reproducible and if the data wasn’t thresholded, we would be able to study the uncertainty in protein interaction networks with a bit more confidence.

References:

[1] Szklarczyk, Damian, et al. “STRING v10: protein–protein interaction networks, integrated over the tree of life.” Nucleic acids research (2014): gku1003

TM-score

 

The similarity between two protein structures can be measured using TM-score (template modelling score). This can be particularly useful when examining the quality of a model, as compared to a target or template structure. One common method of comparing protein structures has been by calculating the root mean squared deviation (RMSD) from the distances of equivalent residues in both structures. An issue with this is that, as all residue pairs are weighted evenly, when the RMSD value is large, it becomes more sensitive to local structure deviation rather than to the global topology. Other established scoring functions, such as GDT-TS (1) and MaxSub (2) rely on finding substructures of the model, where all residues are within a certain threshold distance of the corresponding template residues. However, this threshold distance is subjective and therefore could not be used “as standard” for all proteins. A major disadvantage with all of these methods is that they display power-law dependence with the length of the protein.

TM-score (3) was developed in order to overcome this length dependence. It is a variation of the Levitt-Gerstein (LG) score, which weights shorter distances between corresponding residues more strongly than longer distances. This ensures there is more sensitivity to global topology rather than local structure deviations. TM-score is defined:

where Max is the maximum value after optimal superposition, LN is the length of the native structure, Lr is the length of the aligned residues to the template structure, di is the distance between the ith pair of residues and d0 is a scaling factor. In alternative scoring functions, including MaxSub, d0 is taken to be constant. TM-score uses the below equation to define d0:

which is an approximation of the average distance of corresponding residue pairs of random related proteins. This removes the dependence of TM-score on protein length.

The value of TM-score always lies between (0,1]; evaluations of TM-score distributions have shown that when the TM-score between two structures <0.17, the P–value is close to 1 and the protein structures are indistinguishable from random structure pairs. When the TM-score reaches 0.5, the P-value is vastly reduced and the structures are mostly in the same fold (4). Therefore it is suggested that TM-score may be useful not only in the automated assessment of protein structure predictions, but also to determine similar folds in protein topology classification.

  1. Zemla A, Venclovas Č, Moult J, Fidelis K. Processing and analysis of CASP3 protein structure predictions. Proteins Struct Funct Genet. 1999;37(SUPPL. 3):22–9.
  2. Siew N, Elofsson a, Rychlewski L, Fischer D. MaxSub: an automated measure for the assessment of protein structure prediction quality. Bioinformatics. 2000;16(9):776–85.
  3. Zhang Y, Skolnick J. Scoring function for automated assessment of protein structure template quality. Proteins [Internet]. 2004;57(4):702–10. Available from: http://www.ncbi.nlm.nih.gov/pubmed/15476259
  4. Xu J, Zhang Y. How significant is a protein structure similarity with TM-score = 0.5? Bioinformatics. 2010;26(7):889–95.

Network Representations of Allostery

Allostery is the process by which action at one site, such as the binding of an effector molecule, causes a functional effect at a distant site. Allosteric mechanisms are important for the regulation of cellular processes, altering the activity of a protein, or the whole biosynthetic pathway. Triggers for allosteric action include binding of small molecules, protein-protein interaction, phosphorylation events and modification of disulphide bonds. These triggers can lead to changes in accessibility of the active site, through large or small motions, such as hinge motion between two domains, or the motion of a single side chain.

Figure 1 from

Figure 1 from [1]: Rearrangement of a residue–residue interaction in phosphofructokinase. Left panel: interaction between E241 and H160 of chain A in the inactive state; right: this interaction in the active state. Red circles mark six atoms unique to the residue–residue interface in the I state, green circles mark four atoms unique to the A state, and yellow circles mark three atoms present in both states. In these two residues, there are a total of 19 atoms, so the rearrangement factor R(i,j) = max(6, 4)/19 = 0.32

One way to consider allostery is as signal propagation from one site to another, as a change in residue to residue contacts. Networks provide a way to represent these changes. Daily et al [1] introduce the idea of contact rearrangement networks, constructed from a local comparison of the protein structure with and without molecules bound to the allosteric site. These are referred to as the active and inactive structures respectively. To measure the whether a residue to residue contact is changed between the active and inactive states, the authors use a rearrangement factor (R(i,j)). This is the ratio of atoms which are within a threshold distance (5 angstroms) in only one of the active or inactive states (whichever is greater), to the total number of atoms in the two residues.The rearrangement factor is distributed such that the large majority of residues have low rearrangement factors (as they do not change between the active and inactive state). To consider when a rearrangement is significant the authors use a benchmark set of non allosteric proteins to set a threshold for the rearrangement factor. The residues above this threshold form the contact rearrangement network, which can be analysed to assess whether the allosteric and functional sites are linked by residue to residue contacts. In the paper 5/15 proteins analysed are found to have linked functional and allosteric sites.

Contact rearrangement network

Adaption of Figure 2 from [1]. Contact rearrangement network for phosphofructokinase. Circles in each graph represent protein residues, and red and green squares represent substrate and effector molecules, respectively. Lines connect pairs of residues with R(i,j) ≥ 0.3 and residues in the graph with any ligands which are adjacent (within 5.0 Å) in either structure. All connected components which include at least one substrate or effector molecule are shown.

Collective rigid body domain motion was not initially analysed by these contact rearrangement networks, however a later paper [2], discusses how considering these motions alongside the contact rearrangement networks can lead to a detection of allosteric activity in a greater number of proteins analysed. These contact rearrangement networks provide a way to assess the residues that are likely to be involved in allosteric signal propagation. However this requires a classification of allosteric and non-allosteric proteins, to undertake the thresholding for significance of the change in contacts, as well as multiple structures that have and do not have a allosteric effector molecule bound.

CONTACT

Figure 1 from [3]. (a) X-ray electron density map contoured at 1σ (blue mesh) and 0.3σ (cyan mesh) of cyclophilin A (CYPA) fit with discrete alternative conformations using qFit. Alternative conformations are colored red, orange or yellow, with hydrogen atoms added in green. (b) Visualizing a pathway in CYPA: atoms involved in clashes are shown in spheres scaled to van der Waals radii, and clashes between atoms are highlighted by dotted lines. This pathway originates with the OG atom of Ser99 conformation A (99A) and the CE1 atom of Phe113 conformation B (113B), which clash to 0.8 of their summed van der Waals radii. The pathway progresses from Phe113 to Gln63, and after the movement of Met61 to conformation B introduces no new clashes, the pathway is terminated. A 90° rotation of the final panel is shown to highlight how the final move of Met61 relieves the clash with Gln63. (c) Networks identified by CONTACT are displayed as nodes connected by edges representing contacts that clash and are relieved by alternative conformations. The node number represents the sequence number of the residue. Line thickness between a pair of nodes represents the number of pathways that the corresponding residues are part of. The pathway in b forms part of the red contact network in CYPA. (d) The six contact networks comprising 29% of residues are mapped on the three-dimensional structure of CYPA.

Alternatively, Van den Bedem et al [3]  define contact networks of conformationally coupled residues, in which movement of an alternative conformation of a residue likely influences the conformations of all other residues in the contact network. They utilise qFit, a tool for exploring conformational heterogeneity in a single electron density map of a protein, by fitting alternate conformations to the electron density.  For each conformation of a residue, it assesses whether it is possible to reduce steric clashes with another residue, by changing conformations. If a switch in conformations reduces steric clashes, then a pathway is extend to the neighbours of the residue that is moved. This continued until no new clashes are introduced. Pathways that share common members are considered as conformationally coupled, and grouped into a single contact network. As this technique is suitable for a single structure, it is possible to estimate residues which may be involved in allosteric signalling without prior knowledge of the allosteric binding region.

These techniques show two different ways to locate and annotate local conformational changes in a protein, and determine how they may be linked to one another. Considering whether these, and similar techniques highlight the same allosteric networks within proteins will be important in the integration of many data types and sources to inform the detection of allostery. Furthermore, the ability to compare networks, for example finding common motifs, will be important as the development of techniques such as fragment based drug discovery present crystal structures with many differently bound fragments.

[1] Daily, M. D., Upadhyaya, T. J., & Gray, J. J. (2008). Contact rearrangements form coupled networks from local motions in allosteric proteins. Proteins: Structure, Function and Genetics. http://doi.org/10.1002/prot.21800

[2] Daily, M. D., & Gray, J. J. (2009). Allosteric communication occurs via networks of tertiary and quaternary motions in proteins. PLoS Computational Biology. http://doi.org/10.1371/journal.pcbi.1000293

[3] van den Bedem, H., Bhabha, G., Yang, K., Wright, P. E., & Fraser, J. S. (2013). Automated identification of functional dynamic contact networks from X-ray crystallography. Nature Methods, 10(9), 896–902. http://doi.org/10.1038/nmeth.2592

Addressing the Role of Conformational Diversity in Protein Structure Prediction

For my journal club last week, I chose to look at a recent paper entitled “Addressing the Role of Conformational Diversity in Protein Structure Prediction”, by Palopoli et al [1]. In the study of proteins, structures are incredibly useful tools, offering information about how they carry out their function, and allowing informed decisions to be made in many areas (e.g. drug design). Since the experimental determination is difficult, however, the computational prediction of protein structures has become very important (and a number of us here at OPIG work on this!).

A problem, however, in both experimental structure determination and computational structure prediction, is that proteins are generally treated as static – the output of an X-ray crystallography experiment is a single structure, and in the majority of cases the goal of structure prediction is to produce one model that closely resembles the native structure. The accuracy of structure prediction algorithms is also normally measured by comparing the resulting model to a single, known experimentally-determined structure. The issue here is that proteins are not static – they are constantly moving and may adopt a number of different conformations; the structure observed experimentally is just a snapshot of that motion. The dynamics of a protein may even play an important role in its function; an example is haemoglobin, which after binding to oxygen changes conformation to increase affinity for further binding. It may be more appropriate, then, to represent a protein as an ensemble of structures, and not just one.

Conformational diversity helps the protein haemoglobin carry out its function (the transportation of oxygen in the blood). Haemoglobin has four subunits, each containing a haem group, shown in red. When oxygen binds to this group (blue), a histidine residue moves, shifting the position of an alpha helix. This movement is propagated throughout the entire structure, and increases the affinity for oxygen of the other subunits – binding therefore becomes increasingly easy (this is known as co-operative binding). Gif shown is from the PDB-101 Molecule of the Month series: S. Dutta and D. Goodsell, doi:10.2210/rcsb_pdb/mom_2003_5

How, though, could this be incorporated into protein structure prediction? This is the question being considered by the authors of this paper. They consider conformational diversity by looking at different conformers of the same protein – there are many proteins whose structures have been solved experimentally multiple times, and as such have a number of structures available in the PDB. Information about this is stored in a useful database called CoDNaS [2], which was developed by some of the authors of the paper under discussion. In some cases, there are model (or decoy) structures available for these proteins, generated by various structure prediction algorithms – for example, all models submitted for the CASP experiments [3], where the current accuracy of structure prediction is monitored through blind prediction, are freely available for download. The authors curated a collection of decoy sets for 91 different proteins for which multiple experimental structures are present in the PDB.

As mentioned previously, the accuracy of a model is normally evaluated by measuring its structural similarity to one known (or reference) structure – only one conformer of the protein is considered. The authors show that the model rankings achieved by this are highly dependent on the chosen reference structure. If the possible choices (i.e. the observed conformers) are quite similar the effect is small, but if there is a large difference, then two completely different decoys could be designated as the most accurate depending on which reference structure is used.

The key figure from this paper, in my opinion, is the one shown below. For the two most dissimilar experimentally-observed conformers for each protein in the set, the RMSD of the best decoy in relation to one conformer is plotted against the RMSD of the best decoy when measured against the other:

The straight line on this graph indicates what would be observed if there are decoys in the set that equally represent the two conformers; for example, if the best decoy with reference to conformer 1 has an RMSD of 1 Å, then there is also a decoy that is 1 Å away from conformer 2. Most points are on or near this line – this means that the sets of decoy structures are not biased towards one of the conformers. Therefore, structure prediction algorithms seem to be able to generate models for multiple conformations of proteins, and so the production of an ensemble of models is not an impossible dream. Several obstacles remain, however – although of equal distance to both conformers, the decoys could still be of poor quality; and decoy selection is often inaccurate, and so finding these multiple conformations amongst all others is a challenge.

[1] – Palopoli, N., Monzon, A. M., Parisi, G., and Fornasari, M. S. (2016). Addressing the Role of Conformational Diversity in Protein Structure Prediction. PLoS One, 11, e0154923.

[2] – Monzon, A. M., Juritz, E., Fornasari, S., and Parisi, G. (2013). CoDNaS: a database of conformational diversity in the native state of proteins. Bioinformatics, 29, 2512–2514.

[3] – Moult, J., Pedersen, J. T., Judson, R., and Fidelis, K. (1995). A Large-Scale Experiment to Assess Protein Structure Prediction Methods. Proteins, 23, ii–iv.

The Emerging Disorder-Function Paradigm

It’s rare to find a paper that connects all of the diverse areas of research of OPIG, but “The rules of disorder or why disorder rules” by Gsponer and Babu (2009) is one such paper. Protein folding, protein-protein interaction networks, protein loops (Schlessinger et al., 2007), and drug discovery all play a part in this story. What’s great about this paper is that it gives numerous examples of proteins and the evidence supporting that they are partially or completely unstructured. These are the so-called intrinsically unstructured proteins or IUPs, although more recently they are also being referred to as intrinsically disordered proteins, or IDPs. Intrinsically disordered regions (IDRs) “are polypeptide segments that do not contain sufficient hydrophobic amino acids to mediate co-operative folding” (Babu, 2016).

Such proteins contradict the classic “lock and key” hypothesis of Fischer, and challenge Continue reading

How to Calculate PLIFs Using RDKit and PLIP

Protein-Ligand interaction fingerprints (PLIFs) are becoming more widely used to compare small molecules in the context of a protein target. A fingerprint is a bit vector that is used to represent a small molecule. Fingerprints of molecules can then be compared to determine the similarity between two molecules. Rather than using the features of the ligand to build the fingerprint, a PLIF is based on the interactions between the protein and the small molecule. The conventional method of building a PLIF is that each bit of the bit vector represents a residue in the binding pocket of the protein. The bit is set to 1 if the molecule forms an interaction with the residue, whereas it is set to 0 if it does not.

Constructing a PLIF therefore consists of two parts:

  1. Calculating the interactions formed by a small molecule from the target
  2. Collating this information into a bit vector.

Step 1 can be achieved by using the Protein-Ligand Interaction Profiler (PLIP). PLIP is an easy-to-use tool, that given a pdb file will calculate the interactions between the ligand and protein. This can be done using the online web-tool or alternatively using the command-line tool. Six different interaction types are calculated: hydrophobic, hydrogen-bonds, water-mediated hydrogen bonds, salt bridges, pi-pi and pi-cation. The command-line version outputs an xml report file containing all the information required to construct a PLIF.

Step 2 involves manipulating the output of the report file into a bit vector. RDKit is an amazingly useful Cheminformatics toolkit with great documentation. By reading the PLIF into an RDKit bit vector this allows the vector to be manipulated as an RDKit fingerprint. The fingerprints can then be compared using RDKit functionality very easily, for example, using Tanimoto Similarity.

EXAMPLE:

Let’s take 3 pdb files as an example. Fragment screening data from the SGC is a great sort of data for this analysis, as it contains lots of pdb structures of small hits bound to the same target. The data can be found here. For this example I will use 3 protein-ligand complexes from the BRD1 dataset: BRD1A-m004.pdb, BRD1A-m006.pdb and BRD1A-m009.pdb.

brd1_sgc

1.PLIP First we need to run plip to generate a report file for each protein-ligand complex. This is done using:


 

plipcmd -f BRD1A-m004.pdb -o m004 -x

plipcmd -f BRD1A-m006.pdb -o m006 -x

plipcmd -f BRD1A-m009.pdb -o m009 -x

 


A report file (‘report.xml’) is created for each pdb file within the directory m004, m006 and m009.

2. Get Interactions: Using a python script the results of the report can be collated using the function “generate_plif_lists” (shown below) on each report file. The function takes in the report file name, and the residues already found to be in the binding site (residue_list). “residue_list” must be updated for each molecule to be compared as the residues used to define the binding site can vary betwen each report file. The function then returns the updated “residue_list”, as well as a list of residues found to interact with the ligand: “plif_list_all”.

 


import xml.etree.ElementTree as ET

################################################################################

def generate_plif_lists(report_file, residue_list, lig_ident):

    #uses report.xml from PLIP to return list of interacting residues and update list of residues in binding site

        plif_list_all = []

        tree = ET.parse(report_file)

        root = tree.getroot()

        #list of residue keys that form an interaction

        for binding_site in root.findall('bindingsite'):

                nest = binding_site.find('identifiers')

                lig_code = nest.find('hetid')

                if str(lig_code.text) == str(lig_ident):

                        #get the plifs stuff here

                        nest_residue = binding_site.find('bs_residues')

                        residue_list_tree = nest_residue.findall('bs_residue')

                        for residue in residue_list_tree:

                                res_id = residue.text

                                dict_res_temp = residue.attrib

                                if res_id not in residue_list:

                                        residue_list.append(res_id)

                                if dict_res_temp['contact'] == 'True':

                                        if res_id not in plif_list_all:

                                                plif_list_all.append(res_id)

        return plif_list_all, residue_list

###############################################################################

plif_list_m006, residue_list = generate_plif_lists('m006/report.xml',residue_list, 'LIG')

plif_list_m009, residue_list = generate_plif_lists('m009/report.xml', residue_list, 'LIG')

plif_list_m004, residue_list = generate_plif_lists('m004/report.xml', residue_list, 'LIG')


3. Read Into RDKit: Now we have the list of binding site residues and which residues are interacting with the ligand a PLIF can be generated. This is done using the function shown below (“generate_rdkit_plif”):


from rdkit import Chem,  DataStructs

from rdkit.DataStructs import cDataStructs

################################################################################

def generate_rdkit_plif(residue_list, plif_list_all):

    #generates RDKit plif given list of residues in binding site and list of interacting residues

    plif_rdkit = DataStructs.ExplicitBitVect(len(residue_list), False)

    for index, res in enumerate(residue_list):

        if res in plif_list_all:

            print 'here'

            plif_rdkit.SetBit(index)

        else:

            continue

    return plif_rdkit

#########################################################################

plif_m006 = generate_rdkit_plif(residue_list, plif_list_m006)

plif_m009 = generate_rdkit_plif(residue_list, plif_list_m009)

plif_m004 = generate_rdkit_plif(residue_list, plif_list_m004)


4. Play! These PLIFs can now be compared using RDKit functionality. For example the Tanimoto similarity between the ligands can be computed:


def similarity_plifs(plif_1, plif_2):

    sim = DataStructs.TanimotoSimilarity(plif_1, plif_2)

    print sim

    return sim

###################################################################

print similarity_plifs(plif_m006, plif_m009)

print similarity_plifs(plif_m006, plif_m004)

print similarity_plifs(plif_m009, plif_m004)


The output is: 0.2, 0.5, 0.0.

All files used to generate the PLIFs cound be found here. Happy PLIF-making!

A global genetic interaction network maps a wiring diagram of cellular function

In our last group meeting, I talked about a recent paper which presents a vast amount of genetic interaction data as well as some spatial analysis of the created data. Constanzo et al. used temperature-sensitive mutant alleles to measure the interaction of ~6000 genes in the yeast Saccharomyces cerevisiae [1]. A typical way to analyse such data would be the use of community detection to find groups of genes with similar interaction pattern, see for example [2] for a review. Instead, the authors of this paper created a two-dimensional embedding of the network with a spring-layout, which places nodes close to each other if they show similar interaction pattern.

The network layout is then compared with Gene Ontology by applying a spatial analysis of functional enrichment (SAFE) [3].  Clusters enriched are associated for example with cell polarity, protein degradation, and ribosomal RNA. By filtering the network for different similarities they find a hierarchical organisation of genetic function with small dense modules of pathways or complexes at the bottom and sparse clusters representing different cell compartments at the top.

In this extensive paper, they then go further into detail to quantify gene pleiotropy, predict gene function, and how the interaction structure differs between essential and non-essential genes. They also provide that data online under http://thecellmap.org/costanzo2016/ .

(Left) A network of genetic interaction was embedded into a two-dimensional space using a spring-layout (Right) This embedding was compared with Gene Ontology terms to find regions of spatial enrichment. Image from [1]

(Left) A network of genetic interaction was embedded into a two-dimensional space using a spring-layout (Right) This embedding was compared with Gene Ontology terms to find regions of spatial enrichment.
Image from [1]

References:

[1] Costanzo, Michael, et al. “A global genetic interaction network maps a wiring diagram of cellular function.” Science 353.6306 (2016): aaf1420.

[2] Fortunato, Santo. “Community detection in graphs.” Physics Reports 486.3 (2010): 75-174.

[3] Baryshnikova, Anastasia. “Systematic Functional Annotation and Visualization of Biological Networks.” Cell Systems (2016).

SMEs in Research

Scientific research relies on collaboration, between academics across the world, between big and small groups of people, and between companies and universities. Many people’s first thoughts of companies involved in academia will be large multinational corporations, such as pharmaceutical or aerospace companies. However mutually beneficial relationships exist between smaller companies and academia. Scientific spin out companies, such as Oxford Nanopore Technologies, Theta Technologies or Reinnervate, are one way research is expanded from an idea held by researchers at a university and expanded into a commercial product or service. However, there are many other ways and reasons for scientists collaborating with small companies.

Small and medium sized enterprises (SMEs) are independent businesses with up to 249 people, they represent over 99% of all UK and EU businesses, and 45-50% of turnover and employment. But why would researchers be interested in involvement with SMEs? Access to unique intellectual property and innovation in the SMEs, as well as access to funding targeted at fostering research led projects in SMEs. Innovate UK, support growth by enabling and funding innovative opportunities. SMEs can access this support through Knowledge Transfer Partnerships (KTP), a three-way partnership between a graduate, academic institution and a business. These KTPs can last between 12 months and 3 years, and provide financial support for academics to monitor the graduate’s work. KTP projects lead to an average of 2 papers per project, and are attractive to SMEs as they only fund 33% of the project cost.

Impact of research outside academia is becoming increasingly important to identify as impact case studies form a part of the Research Excellence Framework (REF), making many sources of funding contingent on providing a strong case for the wider impact of the research. Work with innovation focused SMEs can provide a focus for research impact in engagement and economic terms. For example, a REF case study highlighting the impact of spintronics research in the development of non-contact sensors, through a spin out company, Salunda.

Is contacts-based protein-protein affinity prediction the way forward?

The binding affinity of protein interactions is useful information for a range of protein engineering and protein-protein interaction (PPI) network challenges. Obvious applications include the development of therapeutic antibodies to given drug targets or the engineering of novel interfaces for synthetic protein complexes. An accurate model would furthermore allow us to predict a large proportion of affinities in existing PPI networks, and enable the identification of new PPIs, which is critical for our ability to model protein network dynamics effectively.

affinity-prediction-intro

“The design of an ideal scoring function for protein−protein docking that would also predict the binding affinity of a complex is one of the challenges in structural proteomics.” Adapted from Kastritis, Panagiotis L., and Alexandre MJJ Bonvin. Journal of proteome research 9.5 (2010): 2216-2225.

In last week’s paper a new binding-affinity prediction method based on interfacial contact information was described. Contacts have long been used to in docking methods but surprisingly this was the first time that binding affinity was predicted with them. Largely, this was due to the lack of a suitable benchmark data set that contained structural as well as affinity data . In 2011, however, Kastritis et al. presented a curated database of 144 non-redundant protein–protein complexes with experimentally determined Kd (ΔG) as well as x-ray structures.
Using this data set they trained and validated their method, compared it against others and concluded that interfacial contacts `can be considered the best structural property to describe binding strength`. This claim may be true but as we discussed in the meeting there is still some work to do before we take this model an run with it. A number of flags were raised:

  • Classification of experimental methods into reliable and non-reliable is based on what gives the best results with their method. Given that different types of protein complexes are often measured with different methods, some protein classes for which contact-based predictions are less effective may be excluded.
  • Number of parameters for model 6 is problematic without exact AIC information. As Lyuba righlty pointed out, the intercept in model 6 `explodes`. It is no surprise that the correlation improves with more parameters. Despite their AIC analysis, overfitting is still a worry due to the lack of details presented in the paper.

model6-intercept-explosion

  • Comparison against other methods is biased in their favour; their method was trained on the same data set, the others were not. In order to ensure a fair comparison all methods should be trained on the same data set. Of course this is hard to do in practice, but the fact remains that a comparison of methods that has been trained on different data sets will be flawed.

Paper: Vangone, A., Bonvin, A. M. J. J., Alberts, B., Aloy, P., Russell, R., Andrusier, N., … Zhou, Y. (2015). Contacts-based prediction of binding affinity in protein-protein complexes. eLife, 4, e07454. http://doi.org/10.7554/eLife.07454