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Student Training

© Biomatters Ltd 2012

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Table of Contents

Sequences and Alignments ............................................................................. 3

Pairwise Alignment ....................................................................................... 3

Multiple Alignment ...................................................................................... 12

Phylogenetics ............................................................................................. 19

Sequencing .................................................................................................... 22

Full Genome Assembly .............................................................................. 22

Primer Design............................................................................................. 27

Introduction to Biocode LIMS ......................................................................... 32

Components ............................................................................................... 32

Reactions ................................................................................................... 33

Installation of the Biocode LIMS..................................................................... 34

Plugin installation ....................................................................................... 34

Creating your first LIMS database .............................................................. 35

Test that it works ........................................................................................ 37

Remote vs Local ......................................................................................... 39

Data entry and manipulation .......................................................................... 40

What's in a workflow? ................................................................................. 40

Generating plates ....................................................................................... 42

Finding your way around ............................................................................ 65

Some cool things you can do ..................................................................... 67

What about the one-man band? ................................................................. 68

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Sequences and Alignments

Pairwise Alignment

Aligning DNA sequences

For pairwise alignment, you need two sequences which you expect to have a relationship. In the “Pairwise Alignment” folder, select the two pygmy chimpanzee sequences and click the Align/Assemble button on the toolbar and choose Pairwise Align. This will bring up a window that should look like this:

You should reset any changed parameters to their defaults by pressing the

Profiles ( ) button and choosing Reset to Defaults unless it is greyed out in which case you already have the defaults selected.

Click OK and the two sequences will be aligned producing a new document. This will automatically be selected.

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Although you now have an alignment, you also have a dotplot for the two sequences. What you should notice is that the alignment doesn’t look very good in places. There are few gaps and there are regions of high mismatch. This is happening because the default gap penalties are strict and the scoring table is too relaxed. In addition, a pairwise alignment does not deal with sequence inversions.

In the Alignment View locate the first region of mismatch in your alignment. Turn on the consensus sequence and select Highlight Disagreements to make this task easier. Matches will be grey but mismatches will be shaded. Gaps will also be shaded unless you select Ignore gaps which is under the Consensus setting. Mismatching regions will now have a striped effect making them easy to locate. When you have located the first region of mismatches (not gaps), select it and view that region by switching to the Dotplot tab. Turn on the pairwise alignment path option.

This task highlights one of the issues with sequences alignments. The alignment will follow the mathematically optimal path through the two sequences and that does not necessarily reflect what has really happened. It is cheaper for the alignment algorithm to force its way through the mismatches than to take the cost of a high gap penalty.

You should realign these sequences with modified settings. Rather than select the original two sequences, you can just select the previous alignment

document and use Align/Assemble Pairwise Align again as a shortcut.

The gap extension penalty is set to 3 by default and this will limit the distance that a gap can be extended because every gap extension will subtract something from the overall score. Eventually, a gap will become too expensive compared with just accepting a large number of mismatches and this is exactly what happened with the previous alignment. Another problem is that the default scoring table favours mismatches.

To solve these two problems, change the default cost matrix to 93% similarity and change the gap extension penalty to 0, then click OK.

Question 1: What are the coordinates and how do they relate to what you see in that region of the dotplot?

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If you look at the alignment now with the same consensus and highlighting options set, you will see that there is almost no black in the two sequences. If you uncheck ignore gaps it will be clear that the aligner has now used gaps as a way to avoid mismatches. Neither alignment is technically right as there isn’t a correct way to align these two sequences in a pairwise alignment but we’re exploring the data and testing the algorithm.

In the Dotplot there is also an inverted region (you will need to activate the Reverse complement to see this if it isn’t already on). Locate this region on the new alignment by selecting it in the dotplot. This will automatically select the region concerned in the alignment.

In order to align the inverted region you selected in the dotplot (select it again if necessary) and then the same region will be selected in the alignment.

Question 2: What has the alignment algorithm done here?

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Click the Extract button to extract the selected region. You can then re-align as before but be sure to select Automatically determine sequences direction before clicking OK. Geneious will reverse and complement the second sequence selected before doing the alignment. Compare the dotpolot for the extracted region and the re-aligned version. Notice how the diagonal line in the dotplot is now reversed.

Familiarise yourself with the display options that Geneious provides for DNA alignment. Try experimenting with alignment view settings as you will get some interesting effects.

As we have discovered here, the defaults are not necessarily the best for every situation and those that we have used here are also not ideal in every case. When you are ready, move on to the next section.

Aligning Protein Sequences

In this exercise, you will build on your experience with dotplots and DNA alignments to align a pair of proteins. Select the P07990 and ZP_02833023 documents.

If you are not already in the Dotplot view, select it now. You may need to adjust the settings to High Sensitivity / Slow.

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You should see two major regions of homology like this:

Now select Align/Assemble Pairwise Align to perform a standard protein alignment. Select Geneious Alignment and Reset to Defaults if necessary. This will create an alignment document. Turn on the Identity graph, turn off Annotations and turn on both Consensus and Highlighting to make the view clearer.

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These sequences are part of two Type I restriction enzymes and are the DNA sequence specificity domains. Examine the alignment and you will see that the level of identity varies greatly along the length of the alignment. The regions that are conserved are the helical spacers shown in this sketch where the specificity subunit is shaded:

Depending on the family, these helical spacers are highly conserved. However, the Target Recognition Domains are highly variable and this variability is evidence of horizontal transfer. Without the high level of identity in the helical regions, it is unlikely that a decent alignment could be made for these two sequences. The TRD regions can be considered to be aligned purely by chance as the identity is so low. The algorithm has simply computed the cheapest path through this region of mismatch.

In the next example, we have two sequences which are distantly related across their entire length. Select P0A2Q4 and P0ACQ4.

Look a the dotplot and make sure that you have selected the High Sensitivity method. Change the Score Matrix to see what the effect of the different

Question 3: What is the level of pairwise identity between these two sequences and do you think it is a fair representation of their sequence homology? Look in the Statistics tab (%) of the Sequence Viewer options.

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tables is on the dotplot. Choosing a table like BLOSUM30 will show more diagonal matches but this is likely noise. Choose a table which shows a good diagonal for the two sequences without introducing too many short noise lines. Tweaking the Threshold up for instance can reduce the noise while increasing the Window Size will produce longer diagonals.

Align/Assemble Pairwise Align using default settings and examine the result.

The identities here are more evenly spread than in the previous example. This alignment is in the twilight zone for a protein alignment although it has features that suggest it is a good match. Clumping of the identities and the fact that the alignment covers most of the length of the two sequences is a

Question 4: What is the ‘Pairwise % identity’ between these two sequences and is this a fair representation of their homology?

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good sign. Since both these sequences are annotated, you can turn on annotations and you will see that various regions in the two sequences line up well. Manipulate the view using the Zoom functions and other features to observe the details of the alignment and relationship between the sequences.

Question 5: Look at the pattern of identities and regions of mismatch. How do these relate to the annotated domains and regions?

Question 6: There is a region labelled HTH 1 Region in both sequences. How does the pattern of matches and mismatches related to the periodicity in alpha helical structures? Turn on the hydrophobicity graph. Is there a relationship between this and the pattern of mismatches?

Question 7: One of these sequences has secondary structure annotations for the LysR Substrate region. Looking at this alignment and other evidence such as patterns of matches, would you think it is likely that the two sequences share the detailed secondary structure? Justify your observations to support your decision.

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You should now realise that alignments are not a simple process and that alignments do not necessarily represent biological truth. Use them in conjunction with other available information and take care especially when working with distantly related sequences. This will help you understand what is going on with the search parameters and alignments in BLAST searches and also when you start working with multiple alignments.

Question 8: Realign these two sequences using a strict Blosum90 table and Alignment Type: Local Alignment (Smith-Waterman). What has happened to the alignment identity and length? Given what you know about scoring tables and alignment methods, why do you think this has happened?

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Multiple Alignment

A Multiple sequence alignment is a comparison of three or more DNA or amino acid sequences and can be used for many purposes including inferring the presence of ancestral relationships between sequences.

Multiple sequence alignments are more complicated than pairwise alignments because in the ideal case, the alignment would be optimised across all possible combinations of sequences in the alignment. In practice, this isn’t possible so multiple alignment relies on grouping similar sequences together and building out the relationship.

There are several different programs available for multiple alignment in Geneious and they each have sensitivity and performance differences. The most computationally intensive multiple alignment in Geneious is the one provided by the Geneious Aligner.

In the “Multiple Alignment” folder in your training set, select the COXII CDS document. This is a sequence list document. Now click on the

Align/Assemble Multiple Align button and the following dialogue will come up.

This is similar to the alignment dialogue you saw in the Pairwise Alignment section but has some additional features. In this case, turn off the Automatically determine sequences’ direction feature (highlighted) since all sequences are oriented correctly. You can ignore the other features and just click OK.

This aligner is slow and should only be used for relatively small sets of sequences. This set of 51 nucleotide sequences is about the upper limit of sequences that you should consider using with this aligner.

While that alignment is running, you can start another using the included Muscle aligner. Again, use the default settings. Notice how much quicker the

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alignment returns compared with the Geneious aligner. If you open the alignment produced by the Geneious progressive aligner and look at the statistics (%) tab and do the same for the Muscle alignment you should see they are very comparable and yet Muscle finished the job much faster. Most of the time Muscle is a good tool for this.

Now selecting the Muscle alignment, let’s use some of the features of the viewer to make the alignment more interesting. This view has highlighting turned on indicating disagreements with the consensus sequence.

Question 9: Looking at this alignment view, where do you think the sequences are most conserved along their length?

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To get a better idea of the conservation/similarity across the alignment you can switch to the similarity colour scheme. The darker colours indicate the highest conservation.

Interpreting a multiple alignment can be tricky as you can see. Using tools like the identity bar, and the available colour schemes will make it easier.

One of the reasons for producing multiple alignments is to see the effect of mutations on the protein translations of a CDS like this one.

Turn highlighting back on. Now select Translation and choose the Vertebrate Mitochondrial Code. You’ll see another line appear for each sequence which will contain the protein translation of each sequence.

Question 10: Do you still agree with where you thought the highest similarity was from Question 9?

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Zoom in to this view and you’ll see that mutations that change the coding sequence are highlighted. This can be very useful.

You can also turn the translation and highlighting off and switch the colour scheme to Translation which is also useful for indicating non-silent mutations.

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Rather than aligning these sequences as DNA, you may prefer to align as protein. In Geneious, it is simple to translate the sequences to protein by selecting any of your current nucleotide alignments and clicking the Translate button in the toolbar just under the Alignment View tab.

Doing so will produce an unaligned set of amino acid translations. You can’t retain the alignment when you translate like this because the protein alignment may not be optimal.

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If you click Align/Assemble Multiple Align again, this time you’ll be presented with a dialogue which contains protein specific settings such as the Blosum62 cost matrix. Choose Muscle and align the proteins.

Geneious will default to showing a Similarity colour scheme for a protein alignment.

As with DNA alignments, you can use Highlighting to make things more clear. Turn on Highlighting now.

Sometimes, it is useful to perform alignments on DNA as though they were protein. Geneious can do this using the Translation Align tool. Go back to the original COXII CDS nucleotide sequence list and click the Alignment

Question 11: Switching between the DNA and Protein alignments, how does Highlighting behave differently and suggest why?

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button again. This time, select Translation Align and choose Muscle as the engine.

By now you should be quite familiar with how Geneious handles aligning groups of sequences.

Question 12: Compare the alignment produced by the standard DNA sequence alignment with that produced using the Translation Align too. In particular, look at the region around column 150. Are the gaps handled differently and which looks best? What about the identities – which has the best score and does this match which looks best?

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Phylogenetics

Phylogenetics is an important tool for determining evolutionary relationships. In this section, you will be building a tree for the 8 bear sequences which can be found in the “Phylogenetics” folder.

Select all 8 sequences and click Tree. A phylogenetic tree is based on an alignment of all sequences and Geneious will use it’s own pairwise aligner prior to building the tree. Click OK when you’re ready.

Once the tree is built you’ll see a new viewer that looks like this:

So far, you’ve only been using the Geneious Sequence viewer. The Tree Viewer allows you to manipulate the tree in similar ways to the sequence viewer. You can select elements of the tree, change the colours, zoom and change the type of tree displayed. Experiment with this viewer and see what

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it can do. Since this is an unrooted tree, clicking the Enable all layouts for unrooted trees button will let you show other styles of tree.

In addition to the tree view, there is another tab called Distances. Open this.

While this shows you the evolutionary distance between the sequences, it is no substitute for a multiple alignment. Go back to the 8 sequences, reselect

them and this time click Align/Assemble Multiple Align and use Muscle which should produce a good quality quick alignment. The sequence viewer will open this so now you can just click the Tree button as before and Geneious will again produce a tree but it should be much quicker than before since there is now an alignment for the tree to be based on.

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Note that there is no section about the alignment now, and that there are additional options for bootstrapping but don’t use those. Click OK.

Open this tree and it should look much like the previous one. However, there will also be an Alignment View tab which wasn’t there before. This preserves the alignment that the tree was based on and synchronises selection between the tree and alignment views.

The Geneious tree builder is just a basic neighbour joining tree but for publication quality trees you should use one of the other plugins available such as PhyML or MrBayes.

Question 13: Looking at the tree you just produced, which two sequences are most closely related? Check in the Distances view too.

Question 14: Selecting the two most related sequences in the tree will select them in the alignment view. Switch to that view and look at the Statistics tab (%). What is the Pairwise % Identity for these two sequences?

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Sequencing

Full Genome Assembly

With the advent of the current generation of high throughput sequencing technologies it has become possible to rapidly sequence and assemble whole genomes. While de-novo assembly of these technologies is still very time consuming, more rapid mapping of short reads like Illumina data onto a reference genome (so called re-sequencing) is turning up interesting data on SNPs (Single Nucleotide Polymorphisms) for instance.

If you open the “Full Genome Assembly” folder you will find three documents. This is synthetic paired read Illumina data for training purposes but will allow you to see how to use Geneious with real data.

The first step is to select the Forward Reads and Reverse Reads documents. This paired data needs to be processed into a paired reads

document prior to mapping so click Sequence Set Paired Reads… in the menu which will open up the following dialogue:

Set the pairing to the above settings and click OK. Geneious will now set the paired reads and create a new Paired Reads document containing the reads paired together as forward and reverse sequences. Geneious will use this information to assist in placing the reads against the reference sequence because the pairing lets it know how far apart the reads should be to be correctly mapped and this reduces the possible locations where Geneious needs to look in order to correctly place the reads.

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The paired reads document should open in the sequence viewer and look like this:

Now you’ll want to map these reads against the reference genome (in this case, it is the first 1 million bases of E. coli. Select the paired reads document

and the reference genome and click Align/Assemble Map to Reference. Geneious should bring up this dialogue:

Note how Geneious has automatically decided that the longest sequence is the reference. Make sure you choose Do not trim because the data doesn’t need trimming and it would take a long time. You can run with the above values. Click OK and Geneious will map the paired reads against the

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reference sequence. This may take a while so be patient. Once the reads have been mapped, Geneious will open the contig.

At the default zoom level, only the reference sequence, consensus and coverage graphs are visible. You need to zoom in to see the actual reads.

By default, Geneious shows the paired reads linked together with their partner. You can turn this off in the Advanced (cog icon) tab by un-checking the Link paired reads button.

Question 19: Using the default colour scheme, move along the view and look for a SNP (Single Nucleotide Polymorphism). This should appear as an obvious vertical blue line. Where is the first SNP that you can find?

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Finding SNPs manually like this is time consuming so it is better to use a tool to find them for you. Click the Annotate & Predict button and select the Find Variations/SNPs tool. This will bring up the following dialogue:

Click OK and Geneious will scan through the contig and annotate variant sites.

Now that the variant sites are annotated, you can simply click your way along using the < > (inside the red box in the screenshot above) buttons to go to the previous or next annotations. Mousing over the annotation will bring up a tool-tip giving details of the type of variation.

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Annotations are also available in tabular form and this can be very useful when navigating through large data sets. You can also export the annotations as a table that can be imported into a spreadsheet.

In addition to mapping reads, Geneious can also de-novo assemble data from Illumina, SOLiD and 454 machines. With the addition of Geneious Server, users are able to run assemblers and mappers which can’t typically be run on a desktop platform such as Maq, BWA, Bowtie and Velvet. As with the Geneious assembler and mapper used here, these other plugins work in much the same way so should be familiar to you if/when you come to use them.

Question 20: Using the Annotations search and sort functions with the table, how many insertion polymorphisms have a variant frequency greater than 80%? Turn on the polymorphism type column if necessary.

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Primer Design

Geneious provides several operations that work with PCR Primers and DNA or hybridisation probes. PCR Primers and DNA or hybridisation probes can be designed for or tested on existing nucleotide sequences. A PCR product can be extracted from a sequence that has been annotated for both a forward and reverse primer. 5’ extensions consisting of restriction enzymes or arbitrary sequence may also be added to primer documents.

In the “Primer Design” folder, select the DCN gene document. Now select Primers → Design New Primers… which will bring up the following dialogue. Check the product size is set:

Click OK and Geneious will design some compatible primer pairs.

The designed primers are ordered from 1st to 5th with the 1st being the best match for the input parameters. Mouse over any of the primers and Geneious will show a tool tip with details such as melting point, primer-dimer and %GC.

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You can extract these primers by selecting one and clicking the Extract button. This will extract the region annotated as the primer to a new oligonucleotide document. Geneious tracks oligonucleotide sequences as part of the internal primer database so if you want to design with existing primers, these previously designed primers will be offered.

Extract the 1st forward and 1st reverse by selecting each and clicking the Extract button.

Now, delete all primer annotations by clicking the down-arrow for the Primer Bind annotations and choose Delete all primer_bind annotations.

Do the same for the reverse primers and save the document so it no longer has any primers.

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Now you should have an unannotated sequence again. This time select Primers → Test with Saved Primers… and choose the forward and reverse primers you just set as shown in the following dialogue. You can uncheck the product size setting:

This will annotate the primers back onto the sequence as they are compatible:

This is a simple example of the Primer Database. You can specify which primers to use, or you can just let Geneious try all the primers in your local database, or a subfolder and annotate those which are compatible.

With the forward and reverse primers annotated, you can now select Primers → Extract PCR Product which will be the region of sequence bounded by the two primers.

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Try extracting the PCR product now and you should see a result like this:

In this case, the primers were designed within the DCN CDS (hence the ragged ends) but for the purposes of cloning you may want to design primers that will completely enclose the whole CDS. Go back to the DCN gene document and delete the forward and reverse primer annotations and save. You can actually do this just by selecting each annotation and hitting delete since there is only one of each type.

This time, select the DCN CDS annotation because we’re going to design the primers inside this. Again click Primers → Design New Primers… and you should set the following parameters as shown in this dialogue.

Note that the included region is the annotated region of the CDS and the product size is set to the exact length of that CDS in both boxes since you’re going to force Primer3 to design the primers to only be the same length as the CDS. You don’t need to design more than one pair and you need to drop the Tm Min to 52 otherwise there won’t be any suitable primers.

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Now you will see the primers are at both ends of the CDS and if you save and then extract the PCR Product it will be the full sequence of the CDS.

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Introduction to Biocode LIMS The Biocode software comprises everything you need to manage your lab and sequence analysis workflows. These are the key concepts which you should understand before beginning to use the biocode software.

Components

Field database (FIMS)

This stores information relating to field work (e.g. specimen records, collecting events etc). The Biocode plugin does not contain a FIMS database, but rather links to a FIMS database of your choosing.

Lab database (LIMS)

This stores data relating to lab work (e.g. reaction records, plates, etc). It links to data from the Field Database.

Assembler component

This automates the process of assembling your reads, as well as providing useful tools for troubleshooting and analysis. It links to data from both the Field Database and the Lab Database.

Genbank submission

This automates the process of submitting your sequences to Genbank. It combines metadata from the FIMS database with reaction information from the LIMS database, and statistics from the Assembler component to generate a submission.

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Reactions

Plates

You create reactions in plates, which simply groups reactions together in the same way they are in the lab. This grouping represents either physical plates (or strips), or a group of reactions run at the same time. This grouping could be as massively parallel as a 384 well plate, or as small as a single reaction. Both will be covered in this tutorial.

Workflows

Reactions are also linked by workflows. A workflow represents the work done in the lab on one extraction, for a particular locus. It links together a number of reactions carried out at different times, but on the same extraction, and so has information about different PCR reactions tried, what passed and failed etc. Users do not explicitly create workflows - they are created automatically by the LIMS system when plates are saved.

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Installation of the Biocode LIMS

Plugin installation

You will need to download Geneious for Windows, Mac, or Linux, available from the Biomatters website http://www.geneious.com/. To run the Biocode software, you will need Geneious version 5.1 or higher.

Three files, in addition to Geneious, need to be downloaded in order to use the Biocode plugin; the Biocode plugin itself, the Biocode Genbank Submission plugin (which allows you to submit completed contigs to Genbank) and the MySQL Connector (which allows you to connect to FIMS and LIMS databases).

All of these downloads can be retrieved from the Biocode website http://software.mooreabiocode.org/.

For the purpose of this training course, the relevant plugin files have been provided for you. If they are not pre-installed on the machines, the trainer will direct you on the installation of these files.

To install the plugins, open Geneious and drag the plugin onto the Geneious interface. You will see three new icons appear - two in the toolbar and one in the service tree.

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Finally, import the .geneious file that comes with the LIMS training plugins. This file contains a number of sequence files that are used throughout this training manual.

Creating your first LIMS database

You will find the Biocode icon in the Services Panel on the left hand side of the main Geneious window. Right-click (or command-click on Mac) and select Login, and you will be presented with the Biocode Login screen, and then click the Add button. The next step is to set up a connection to a field management (FIMS) database, and a Laboratory Management (LIMS) database.

FIMS configuration

The Biocode LIMS supports connections to many types of FIMS databases, from simple one-user systems through to commercial packages. Throughout this course, we will be using the simplest case: an Excel spreadsheet containing our FIMS information.

Choose Excel from the Field Database Connection dropdown, select the Browse button, and navigate to the file example_FIMS.xls which is located in the same folder as this document.

Now we need to give the plugin some information about how our Excel file is structured.

Select the Tissue, Specimen, Plate, and Well fields from the dropdowns, as shown in the illustration below. You may press the Autodetect button to fill in the taxonomy values.

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LIMS configuration

The Biocode Plugin can connect to remote LIMS databases, supporting many users, and can also connect to a built-in LIMS database more suitable for tracking a single user’s lab workflow. For this example, we will be using a built-in LIMS database.

Select Built-in MySQL database from the LIMS Connection dropdown, and click Add Database. Enter a name for your database, and click OK.

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That’s it! Click OK on the Biocode Connections dialog to connect to your new LIMS database.

Test that it works

The easiest way to ensure that you’ve successfully set up your LIMS database (and it’s connection to your existing FIMS database) is to try and search for existing data and to add a simple reaction plate.

The sample FIMS database has taxonomic metadata attached to the tissue samples. To see if this data can be accessed, try searching for Gastropoda and examine the documents that are returned.

What kind of documents do you see returned in the document table?

Shown is a collection of tissue samples that have the keyword _Gastropoda_ in one of the searchable fields. In the case of these documents, the class field contains the term.

Note the various fields that are visible in the document table list.

Later, users will be shown advanced search techniques to limit the search to particular fields.

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The above exercise tested that information could be read from the FIMS data source. As we have created a new, custom LIMS database we do not need to test that we can write to the database at this time. This is because the database was created at the time of login. The fact that this creation step did not give an error message means that the database now exists and is writable by the Biocode plugin software.

We will start entering data into the LIMS system in the next session.

Thumbs up!

Congratulations! You have set up your first Biocode LIMS database!

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Remote vs Local

It is worthwhile discussing the other connection options at this point. You can connect to one of four types of FIMS, and two types of LIMS databases

FIMS

● TAPIR FIMS - This is most useful for groups who want to store their lab information in a single FIMS, or for collaborative projects. The TAPIR FIMS will read tissue data from a TAPIR server. For more information see http://www.tdwg.org/standards/449/

The following table-based FIMS connections allow you to specify your own data structure:

● MySQL FIMS - This is most useful for groups of users who have an existing FIMS system backed by a MySQL database, but do not want to set up a TAPIR server. Geneious will read the data directly from the database.

● Google Fusion Tables FIMS - This is most useful for groups of users who want to access their FIMS data remotely, and do not already have a compatible way of storing it.

● EXCEL FIMS - This is most useful for single users, or people who cannot set up a TAPIR server. The EXCEL FIMS will read tissue data in from an excel sheet.

LIMS

● Remote LIMS - The plugin will store your lab workflows on a remote MySQL server. This is the best option for labs, or collaborative projects.

● Local LIMS - The plugin will store your lab workflows on your local machine. This is intended for single users.

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Data entry and manipulation

What's in a workflow?

Workflows are an integral part of the LIMS system, so it is important that you understand how they work before entering any data.

Workflows represent the path that an extraction takes through the lab, for a particular locus. A workflow is limited to a single extraction, but can have any number of other types of reactions. The only restriction is that each reaction in a workflow must have the same locus.

Workflows are useful for tracking the progress of work, and for troubleshooting problems in the lab. Information from workflows is also taken by the assembly module to aid in the assembly and analysis of your sequences, so it is important that they are set up correctly. The diagram below describes an example workflow.

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You should note the following points:

● One extraction can belong to many workflows (as extractions are often used as stock for many reactions)

● You can have any number of failed reactions in a workflow, and you can have any number of passed reactions in a workflow. The passed or failed status of a workflow for a given reaction type is taken from the most recent reaction of that type.

● While each workflow can only contain reactions of a single locus, each locus can have any number of workflows (useful if multiple people are working independently on the same locus for the same extraction).

Workflows are created with reactions. Any reaction (apart from extractions) that has an empty workflow field when saved, will have a new workflow created for it. That means that it is particularly important that you fill in the workflow field correctly for all reactions that you save. Fortunately, this is easily accomplished in the bulk-editor. Clicking autodetect workflows in the tools dropdown will automatically fill in the workflow field for any reactions that have an available workflow (i.e. one with a matching extraction and locus).

If no matching workflow exists, the workflow field will remain blank, and a new workflow will be created when you save the reaction.

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Generating plates

As covered at the end of the last section, your new LIMS database contains tissue information (from the EXCEL sheet you used for your FIMS), but no laboratory information. Try searching for Gastropoda, and examine the documents that are returned. We are going to create an extraction plate from these tissue records.

Extraction plate

Click New Reaction in the Geneious Toolbar to open the reactions dialog.

Choose Extraction as the reaction type, and choose 96 well as the plate size. Click OK, and you will see a plate editor showing a blank plate. The Biocode Plugin has a bulk-editor to enable you to quickly enter reaction id’s into the plate. Click Bulk Edit in the plate editor toolbar to open the bulk editing dialog. The bulk editing dialog contains a table, where each column represents a field on the reaction, and each row represents a well in the plate. You can paste in values from the clipboard, but in this case we are going to copy tissue id’s directly from the FIMS database.

Select Tools, then select Get Tissue ID’s from Archive Plate. Since our FIMS database is organised into plates, we can automatically copy the tissue id’s into the correct well. The sample FIMS database contains information for one plate. Enter ‘Plate_m037’ (without the commas) into the dialog, and click OK.

You will see the Tissue ID column filled with data. Select Tools, then select Generate Extraction Id’s to automatically generate id’s for your new extractions.

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Click OK to return to the extraction plate, and you will see the wells filled with values.

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Select Edit All Wells from the plate editor toolbar to enter lab information about your extractions (such as volume and concentration), and click OK when you’re done. Set the date of creation to be a day earlier from today.

Finally, to save your extractions, enter a name for your plate (in the top left corner of the editor), and click OK.

Note: For the purpose of this exercise, we are using ‘myExtraction’ as the name of our extraction plate. You can use your own name, but remember to replace all instances of myExtraction with the name you chose for the remainder of the tutorial.

To check that everything worked correctly, you can search for your plate by typing its name into the search box.

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PCR plate

Geneious can use an existing plate as a guide for creating a new plate, so that the genetic material in each well of the new plate corresponds to the genetic material in the existing plate. To do this an existing plate document is selected in Geneious and a new plate is generated using the selected plate as a template.

If the reaction types are the same (for example, you're creating a PCR plate from a PCR plate), then all reaction parameters will be copied to the new plate. If the reaction type is different (for example, an Extraction to a PCR plate), then only the extraction id's will be copied across.

We will use this method to generate a PCR plate from the extraction plate we created in the previous section. To find the extraction plate we just created, search for ‘myExtraction’ and check the plate checkbox to bring up the plate we just made.

Select the extraction plate that shows up in the results. Click new reaction. In the dialog that shows, ensure that Create plate from existing document is checked and choose PCR as the plate type.

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This will produce a skeleton plate template to define your PCR reactions on the samples from the extraction plate.

The default label for a PCR reaction plate is to show the primers used for the PCR and the reaction cocktail used. One of the first things you will notice from the generated plate is that it is devoid of useful label information, currently having no primers or reaction cocktail selected for the samples on this plate.

To add reaction information to this plate, selecting ‘edit all wells’ from the top right of the screen will open up the bulk editing screen.

The first thing we will want to change is the reaction date, setting it again to yesterday (as was done in the extraction plate step).

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There are several fields here that need to be filled out before the PCR plate can be saved. You should make sure there is an entry for the following fields before you leave this window:

● Locus ● Forward Primer ● Reverse Primer ● Reaction cocktail

The first, Locus, is a simple text field. You can either select from one of the common barcoding entries from the dropdown list, or give your own locus name should you be working on a novel locus, or have an internal naming convention used for loci in your research group.

The other three compulsory fields are more complex. The primer fields are populated by selecting oligo-nucleotides from your local Geneious database.

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The reaction cocktail is selected from a list of created cocktails in your LIMS database. As this is your first reaction plate in your local database this cocktail list will be blank, a new cocktail will need to be defined.

Primers

Firstly, let us populate the primer fields. Click the Choose... button next to the primer fields. This is display a list of all Primer documents in your local Geneious database. Simply select the appropriate primers for your reaction and press Select. For this tutorial use M13_F-39dgLCO1490 and M13_R-dgHCO2198 to populate the forward and reverse primer fields, respectively.

Cocktails

Cocktails are a recipe for the ingredients that will go into a reaction (excluding the primer). You can choose from a list of existing cocktails, or create your own.

To create your own cocktail, click “Edit Cocktails”, then click new in the dialog, and enter the volumes and concentrations in the fields provided.

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Note: you cannot modify cocktails once they are created. However, you can create a copy of an existing cocktail by selecting it in the view, and then clicking “Add”. This will create a new cocktail for editing, with the field contents copied from the previously selected cocktail.

As opposed to being a limitation, the inability to edit a previously created cocktail is a purpose-designed feature. Why do you think this is the case?

The reason to disallow editing of cocktails is for the purpose of record-keeping and reporting. Reports can be created that trace the usage of various reagents. Furthermore, a workflow shows the history of what has been produced from a sample.

Editing the cocktail would in essence change the history of all workflows using the cocktail, resulting in potentially incorrect and misleading audit trails.

Once you have entered data into the four required fields, press OK on the Edit Wells dialog to propagate the changes to the PCR plate. You will notice that

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the previously empty label fields have now been filled with the primer and cocktail information.

The workflow field was left blank when filling out the details of the reaction. Why was this and what effect will it have on the LIMS data?

Workflows are created with reactions. Any reaction (apart from extractions) that has an empty workflow field when saved, will have a new workflow created for it. That means that it is particularly important that you fill in the workflow field correctly for all reactions that you save.

Fortunately, this is easily accomplished in the bulk-editor. Clicking autodetect workflows in the tools dropdown will automatically fill in the workflow field for any reactions that have an available workflow (i.e. one with a matching extraction and locus). If you do want to create new workflows for your reactions, just leave the field blank.

Thermocycles

You can choose a thermocycle for PCR and cycle sequencing plates from the drop down list in the plate editor toolbar, or create your own using the editor.

The thermocycle editor comes in two parts – a cycle editor, and a state editor.

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Start by creating the cycles you want by clicking the + button below the thermocycle diagram. Enter the number of times that you want the cycle to repeat in the selector, and click the “Edit Cycle” button to add states to your cycle.

You can similarly click the + button in the states dialog to add states to your cycle, and you can give each state a temperature and a time. Check the “Infinite” checkbox to make the state last for an infinite amount of time.

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At this time you cannot add advanced features such as temperature gradients to your thermocycles, but there is a space for notes where you can list these.

Try making a thermocycle with the below characteristics for your PCR reaction plate and give it a sensible name. ‘default CO1 PCR cycle’ might be a good one:

You now have a PCR plate set up with primers, a cocktail and thermocycle defined. You have also given the plate some distinguishing information in the various plate fields (more on how to access and use these later).

Taking it to the lab

Once you have defined the various fields for your reaction and attached the relevant primers, cocktail and thermocycle you are ready to take it all in to the lab.

Plate documents can be printed to create a template used to fill the actual plate in the lab. It will print out a diagram of the plate, a list of recipes for all of the cocktails on the plate (listing enough quantities to make cocktail solutions for the entire plate), the thermocycle (except for extraction plates), and a legend if you are printing the plate with color.

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Select the fields that you want to be displayed on the plate using the Display Options before printing, and they will be displayed in your plate printout.

The plate editor can display any number of fields from your FIMS or LIMS database. Click the "Display Options" button in the toolbar to open the display dialog.

The split-pane at the center of the dialog allows you to choose which fields to display on your wells. The available fields are shown in the left hand pane. Select the fields you want to display, and click the right arrow to move them to the right hand pane. Select fields in the right hand pane and click the left arrow to return them to the right hand panel, and stop them from being displayed in the wells. Once you have chosen your fields to display, you can choose their order using the up/down arrows on the right hand side of the dialog. The fields will appear in the wells in the order you choose here.

The bottom part of the dialog controls the well coloring. Wells are colored by one field, with each different value for that field being given a different color.

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Choose the field you are interested in, and all possible values will be displayed below it. You can change the color of any value by clicking it.

You can save any settings you make here as a template by clicking the "Select a template" button at the top of the dialog, and clicking "Create template". Click the "Save as Default" button to make that template the default (separate defaults are stored for Extraction, PCR, and Sequencing plates).

Below is a PCR plate report showing a plate of reactions that use two different cocktails. The plate has been coloured by cocktail, and the cocktail has been made the primary field for the display. Try to replicate this display view using your myPCR plate. Save this new display as a display option template (but DON’T save it as the default view).

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GEL images

You can attach GEL images to all types of plates. Click the “Add/Remove GEL Image” button in the plate editor toolbar, and click “Add”, then browse to the image file on your hard disk. In the case of this tutorial, browse to the location of your tutorial files on your computer. Select the image file called ‘gelImage.jpg’.

Note: In this example we used a JPEG (.jpg) file. Geneious will accept images in multiple formats, including JPEG, GIF, PNG, and TIFF formats.

You can split a GEL image, and attach the segments to each well by clicking the “Split GEL” button (located above the GEL image in the gel viewer window).

GEL images are split into wells using a dragable grid, where each cell in the grid represents a well.

Ensure that the options are set as follows:

Number of Rows: 8

Number of Columns: 12

Start: A

Start: 1

Automatically score plate: Yes

Threshold: 27

Once you have set the above, Rotate the image until the gel columns look level by using the slider below the zoom controls, then drag your mouse on the image to create the grid. If you misplace the grid, you can start again by dragging the mouse. Click ok when done. You can also add notes to each GEL image.

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Once you have scored the gel, press OK again in the gel adding dialog. You will find that the various wells will be coloured either red or green. Looking at the legend will show that green represents a passed reaction and red a failed. These values were set by the automated scoring function of the gel splitting operation.

If you set the previous setting correctly, the plate should show wells A3 and F4 as failed reactions.

The automated scoring function is not perfect and it is worthwhile manually verifying the results. The best way to do this is view the plate with the gel images superimposed, whilst still colour-coded for the reaction state.

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To do this, go to Display Options. Remove the current displayed tabs from the list of Selected fields, and add the GELImage field so that it is the only one showing. In the bottom section, ensure Color wells based on is set to ‘Reaction state’. If you prefer labels instead of colour coding due to printing or other constraints, you should also add the Reaction state as a field. See the below image for what the display options panel should look like.

Once the display options have been set as in the above panel, you should be presented with a plate viewer that looks like this:

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You will notice by way of brief visual inspection that the automated scoring was not perfect: There are some reactions of dubious success in this plate. Looking at the plate, we might decide that A9 is also a failed reaction. Double-clicking the well will bring up the Well editing dialog for the A9 well. In this dialog, change the Reaction state field to show ‘failed’. Pressing OK will take you back to the plate view which should now additionally show the A9 well as being a failed reaction. A fast way to do this for multiple wells is to hold down the ctrl key (the command key on mac), and click on all the wells you want to fail. Then click Edit Selected Wells, and change the reaction status to failed.

Once you have done this, you should save the changes to the plate.

Sequencing plate

The PCR plates have been processed and you now know which of the reactions have been successful. It is time to create a sequencing plate for the passed reactions (re-running failed reactions will be covered later, for those wondering what will happen to those failed PCRs).

Search for your created PCR plate so that it shows in the document table view. Select the document and press the ‘New reaction’ button from the toolbar. We are going to create a new plate from an existing one in a similar fashion to how we created the PCR plate.

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This time however, we wish to only create wells for our passed PCR reactions. To do this, enable the Copy only Passed reactions tickbox. Ensure that you are creating a plate from the existing document and that you are creating a new Cycle Sequencing plate.

This will create a new plate with wells for the sequencing of the passed PCR reactions. Notice the blank well positions marking where the failed PCR reactions were in the template plate.

Once you have your plate template, press ‘Edit all wells’ to populate the various fields for the sequencing. Many of the fields have been copied over from the PCR plate (and the extraction plate before that one). You will need to fill out the primer and cocktail fields.

This plate is going to be for sequencing the forward strand. Under primer, ensure that the direction is set to forward and choose the forward primer from the PCR step (M13_F-39dgLCO1490) by pressing the ‘Choose’ button.

You will also want to create a cocktail for the sequencing reactions.

Create a cocktail with the following settings:

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Press OK once you’ve edited the wells and have settings similar to those in the above screenshot.

Similar to the steps from the PCR plate, you will also need to create a thermocycle for the sequencing plate.

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Using the process you learnt in the PCR plate section, create a sequencing thermocycle that conform to the below thermocycle diagram:

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Once you have created this thermocycle, give the plate a meaningful name (myForwardSeq) and save the results.

Repeat the above steps to create a plate to sequence the reverse strand, using the thermocycle and cocktail you just created.

Are there any meaningful display options you could create at this point?

Adding traces (getting back from the lab)

Once you’ve performed the sequencing tasks in the laboratory and produced trace files from your sequencing reactions, it is time to revisit your sequencing plate and link it with the trace data you’ve generated.

Before we can do this, there are some preparatory steps that need to be taken with your data. You will not have to do this for the tutorial (they have already been prepared) but it is good to have a record of the required steps.

Traces for a plate are added at the folder level. This means that the forward plate and reverse plate must have their trace files in separate folders. These folders must contain no data other than trace files for the plate. Secondly, each trace file must have a reference to the source well (i.e. A01, H12, etc).

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This can be part of a larger filename, as long as it is delimited using some kind of control character (e.g. an underscore or a space character).

The structure should be something like this:

You will find the previously prepared data for this tutorial under the traceData folder in your tutorial files.

Search for and select the first (forward) direction sequencing plate created earlier called myForwardPlate. With this selected, press the ‘Bulk add traces’ button in the plate view toolbar.

Press Browse and navigate to the location of the traceData folder and select the folder called ‘forward’.

In the fields below the file selector, use the well number field as the match. The match field specifies which field in the LIMS the importer will look for in the imported filename to match the trace file with the correct well. In most cases you will want to use the well number.

For our sample data, match the 2nd part of the name, separated by the underscore character.

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You should end up with a dialog similar to that below:

Given the example filenames given in the folder image on the previous page, what match filter would work with those files?

Match well number

Match 3rd part of name, separated by Underscore

To check if the traces have added correctly, double-click one of the wells to bring up the Edit well(s) dialog. There is a button in this view called ‘Add/edit traces’. Pressing this button will bring up a sequence viewer representation of the trace, along with its base calling and associated quality.

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Accept these open dialogs to go back to the main Geneious window.

Finding your way around

Biocode searches return four types of documents:

1 Tissue sample documents – Each of these represents a tissue sample in the field database. Tissue documents contain collection information, and optionally taxonomy and photographs.

2 Plate documents – These represent a plate in the lab database, and contain a diagram of the wells, as well as the plate’s thermocycle and attached GEL images if available.

3 Workflow documents – These contain a linked set of reactions performed on an extraction.

4 Sequence documents - sequences entered into the LIMS when traces/assemblies are marked as pass/fail.

You can perform either a basic search or an advanced search. Basic searches are performed by entering text into the search box, and return all

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documents which have a field with a similar value to the text entered. You can restrict searches to particular types of documents by unchecking some of the checkboxes to the right of the search box.

Advanced searches explicitly search against particular fields. They are performed by clicking the “More Options” button. Click the + and – button to add and remove fields from the search. Choose the fields you want to search using the leftmost dropdown, and choose the search condition using the dropdown box to its right.

Create searches for the following scenarios:

1. All work done after the beginning of May

Last Modified (LIMS) | Greater Than | 01-May-2010

2. All plates which have the phrase “my" somewhere in their name

Plate Name (LIMS) | Contains | “my”, only plate checkbox checked

3. All COI workflows and plates

Locus | Contains | “COI”, only workflow and plate checkboxes checked

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Some cool things you can do

Converting between 96 and 384 well plates

It is possible to create a new 384 well plate from a group of 96 well plates, and to create a group of 96 well plates from a 384 well plate. Each 96 well plate corresponds to one quadrant in the 384 well plate (see below).

To create a 384 well plate, select up to four 96 well plates in Geneious, and click “New Reaction”. Select the “Create plate from existing document” checkbox, and choose “384 well plate”. A panel will appear at the bottom of the dialog which will allow you to choose which quadrant each 96 well plate corresponds to.

Cherry picking

The Cherry Picking function allows you to select reactions from one or more plates, based on criteria that you specify (e.g. failed reactions, or genes of a particular taxonomy). You can use these selected reactions to create a new plate (or plates), or have them returned to you as a list.

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To perform Cherry Picking, select the plates containing the reactions you want to pick, and click on Cherry Picking in the Biocode toolbar menu. Choose your destination, and then choose the criteria to select your reactions.

What about the one-man band?

The Biocode LIMS software, as seen above, is fully capable of scaling into a large-scale operation. In the situation of a smaller laboratory environment, you may be working with sample tubes as opposed to an array of wells. In the Geneious LIMS package, this works in a very similar fashion, with a single reaction tube being considered a single-well plate.

To highlight the differences, lets take a sample through from extraction to sequencing. We will have a look at the assembly stages in the Data analysis section.

In the last section we were creating plates using information from archived plates in the LIMS. With a single-reaction approach we will work from a tissue sample point of view.

Extraction

Search for ‘MBIO’ in the LIMS search dialog with only the Tissues checkbox enabled. This will show up a number of tissue samples from the LIMS. Making sure the tissue_id column is enabled and visible, note down a tissue id of interest. For this exercise, we are interested in MBIO19229.1.

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Ensuring no tissue is selected in the document table, press the New Reaction button in the toolbar. We are making an Extraction reaction, with one individual reaction, as per the below image:

This will result in a new blank Extraction plate of one well:

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To get started with this plate. Press the ‘Bulk edit’ button. In the Tissue Sample Id field, put in the tissue id we identified earlier, “MBIO19229.1”. Once you have done this, go to Tools > Generate Extraction IDs.

You will note that the extraction ID ends in the digit 2. Can you explain why this might be the case?

This is because it is the second extraction from this tissue sample. The auto-extraction ID generator adds an incremental number after the sample Id. this can give a rough indication on the number of times this sample has been processed

Once you have edited the above core details, you can double-click the A1 well to fill out any remaining details about the extraction, including the name of the technician that performed the extraction.

Once you have done this, save the plate with the name ‘singleExtraction’.

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Remember, the LIMS can be a multi-user environment, so having the ability to trace who is performing even single reactions can be useful for reporting purposes.

PCR

The PCR stage will be almost identical to the PCR stage from the multi-well plate.

Try to generate a single PCR reaction, using the ‘singleExtraction’ reaction as a template. Possible points of difference will be outlined below.

One point of difference may be how you handle PCR images for a single reaction. When adding the image, you may have a fuller GEL image as below, complete with scaling bar and negative control.

We have an image such as this for the purposes of this tutorial, using the GEL images function, add the singleGelImage.jpg file to the PCR reaction plate.

Given that you are working with a single reaction, manual verification is more than sufficient. Despite this, you may wish to use the split gel function to highlight the PCR band of interest. The section you identify using the split gel function can be used as a display field on the summary page.

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Once you’ve identified the PCR band region of the image, adapt the display options to show the section of the PCR image on the summary display. See below for an example screenshot.

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Sequencing plate

The generation of sequencing single reaction plates works, again, in the same way as it did in the multi-well scenario. The way traces are added however will be different.

Using the Sequencing section from the multi-plate sub-section as a guide, create two single reaction plates from the single PCR plate - one using the forward primer and the other using the reverse. Name them singleForwardSequencing and singleReverseSequencing respectively.

Once both plates have been created, use the search function to find them both again.

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Select the forward sequencing plate. Double-click the well to bring up the well editing dialog. Under this page you will find a button to Add/edit traces. Press this button to bring up the below dialog.

Using the file dialog, browse to the tutorial files. Upload the file “9823744_F.ab1” from the singleTraces directory.

Once you have done the above and saved the plate. Do it again for the reverse plate, this time using the “9823744_R.ab1” trace file.