How should I ship plasmids?

How should I ship plasmids?

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I shipped 10 µL of my vector miniprep to a collaborator in a 1.5 mL eppendorf parafilmed shut and stuffed into a 50 mL conical with some paper-towel padding. However, something happened on the way and there was nothing (no liquid) in the tube when it arrived. They didn't make any comments about the microcentrifuge tube popping open or broken parafilm, so nothing crazy happened but something did.

What's the most reliable way to ship plasmids?


  • the 10 uL of plasmid miniprep may have been splattered in the cap of the tube (AnnaF)
  • the eppendorf tube may have depressurized during air shipment and allowed the 10 uL to escape and evaporate
  • solution: try air-drying or blotting (Jonas) your minipreps prior to air shipment


As AnnaF wrote, the 10 uL of your plasmid could have been hidden in the cap or dispersed around the tube, making appear empty. You should check with your collaborators to be sure they centrifuged it.

According to a fedex document on shipping perishables (pdf) and a paper measuring the temp and pressure of air shipments (pdf), fedex and ups air shipments may experience low pressure environments around 0.56 - 0.74 atmospheres (atm). At these relatively low pressures, perhaps an eppendorf tube sealed at 1 atm might breach. The papers also note that ground shipments that pass over the rockies (i.e. in Colorado) may experience ~ 0.64 atm.

So perhaps your 1.5 ml eppendorf tube depressurized during the shipment?

It would be interesting to do some tests on the pressure-worthiness of eppendorf tubes.

Regarding the original question, in 2007 I prepared and shipped (fedex) a library of thousands of minipreps to hundreds of users. 1uL of miniprep was dispensed into wells of 384-well plates and airdried, then sealed with aluminum, then mailed. Users rehydrate a well with 10 uL of water. Generally it works.

A quite safe way of shipping plasmids is to put them on filter paper (see protocol and send a letter. Much cheaper.

Did they try to centrifuge the tube when it got there to push all the liquid to the bottom? I know that especially when working with such little amounts that even shaking it up a little can disperse the contents all over the tube. We have received plasmids from other labs before. Generally speaking the plasmids are sent in Screw-cap microcentrifuge tubes inside of some sort of canister. This is then packaged in a little foam cooler with dry ice to keep everything cold.

Since tubes can be crushed in the mail, the safest way to ship plasmids is to drip a few uL into a filter paper, and then wrap it up to seal it with parafilm, and fill out a detailed form about its content.

Good luck.

We have gotten into similar situations when other labs have sent us plasmids (or when we have taken out ancient tubes out from storage boxes at -20), and have since adopted the filter paper method. Another point to note is that you can just add, say, 10 μL of water to the "empty" tube (Mac) and use 1 μL for a transformation. It has always worked for us. DNA is seemingly invisible!

I will also strongly recommend blotting a microgram of your plasmid DNA on a piece of filter paper (the filter paper is important for extraction on the receiving end). Also, it is very very helpful if you clearly indicate the amount of DNA (approximately) and circle the blotted DNA in pen so there is no ambiguity about cutting out the circle of filter paper containing the DNA.

On the receiving end, you should always freeze the paper right away or elute the DNA off of the paper in TE buffer or water and then freeze. Failure to do so will eventually lead to DNAase degradation of the plasmid DNA and then you'll have to wait for another shipment.


David P. Clark , . Michelle R. McGehee , in Molecular Biology (Third Edition) , 2019

2 General Properties of Plasmids

Plasmids are usually circular molecules of DNA, although occasionally, plasmids that are linear or made of RNA exist. They may be found as single or multiple copies and may carry from half a dozen to several hundred genes. Plasmids can only multiply inside a host cell. Most plasmids inhabit bacteria, and indeed around 50% of bacteria found in the wild contain one or more plasmids. Plasmids are also found in higher organisms such as yeast and fungi. The 2 micron circle of yeast (discussed later) is a well-known example that has been modified for use as a cloning vector.

Most plasmids are circular, made of DNA, and much smaller than chromosomes.

The copy number is the number of copies of the plasmid in each bacterial cell. For most plasmids, it is 1 or 2 copies per chromosome, but it may be as many as 50 or more for certain small plasmids such as the ColE plasmids. The number of copies influences the strength of plasmid-borne characteristics, especially antibiotic resistance. The more copies of the plasmid per cell, the more copies there will be of the antibiotic resistance genes, and therefore, the higher the resulting level of antibiotic resistance.

The size of plasmids varies enormously. The F-plasmid of Escherichia coli is fairly average in this respect and is about 1% the size of the E. coli chromosome. Most multicopy plasmids are much smaller (ColE plasmids are about 10% the size of the F-plasmid). Very large plasmids, up to 10% of the size of a chromosome, are sometimes found, but they are difficult to work with and few have been properly characterized (see Box 23.02 ).

When the genome of the Gram-negative bacterium Vibrio cholerae, the causative agent of cholera, was sequenced, it was found to consist of two circular chromosomes of 2,961,146 and 1,072,314 bp. Together, this totals approximately 4 million base pairs and encodes about 3900 proteins—about the same amount of genetic information as E. coli. Many genes that appear to have origins outside the enteric bacteria, as deduced from their different base composition, were found on the smaller chromosome. Many of these genes lack homology to characterized genes and are of unknown function. The smaller chromosome also carries an integron gene capture system (see Chapter 25 : Mobile DNA) and hosts “addiction” genes that are typically found on plasmids (discussed later). Furthermore, the smaller chromosome replicates by a different mechanism from the large chromosome. In fact, the smaller chromosome shares a replication system with a family of widely distributed plasmids. It seems likely that the smaller chromosome originated as a plasmid that has grown to its present size by accumulating genes from assorted external sources. Perhaps it is better to regard the smaller chromosome as a “megaplasmid.” Genome-sequence data suggest that some 10% of bacteria carry such megaplasmids, although the size varies considerably. In most of these cases, the larger chromosome carries almost all of the genes needed for vital cell functions such as protein, RNA, and DNA synthesis. In a few cases such megaplasmids can transfer themselves to related bacteria by conjugation.

Some plasmids are present in one or two copies per cell, whereas others occur in multiple copies.

Plasmids carry genes for managing their own lifecycles and some plasmids carry genes that affect the properties of the host cell. These properties vary greatly from plasmid to plasmid, the best known being resistance to various antibiotics. Cryptic plasmids are those that confer no identifiable phenotype on the host cell. Cryptic plasmids presumably carry genes whose characteristics are still unknown. Plasmids that are modified for different purposes are used in molecular biology research and are often used to carry genes during genetic engineering.

The host range of plasmids varies widely. Some plasmids are restricted to a few closely related bacteria for example, the F-plasmid only inhabits E. coli and related enteric bacteria like Shigella and Salmonella. Others have a wide host range for example, plasmids of the P-family can live in hundreds of different species of bacteria. Although “P” is now usually regarded as standing for “promiscuous,” due to their unusually wide host range, these plasmids were originally named after Pseudomonas, the bacterium in which they were discovered. They are often responsible for resistance to multiple antibiotics, including penicillins.

Certain plasmids can move themselves from one bacterial cell to another, a property known as transferability . Many medium-sized plasmids, such as the F-type and P-type plasmids, can do this and are referred to as Tra + (transfer-positive). Since plasmid transfer requires over 30 genes, only medium or large plasmids possess this ability. Very small plasmids, such as the ColE plasmids, simply do not have enough DNA to accommodate the genes needed. Nonetheless, many small plasmids, including the ColE plasmids have mobilizability , meaning they can be mobilized by self-transferable plasmids [i.e., they are Mob + (mobilization-positive)]. However, not all transfer-negative plasmids can be mobilized. Some transferable plasmids (e.g., the F-plasmid) can also mobilize chromosomal genes. It was this observation that allowed the original development of bacterial genetics using E. coli. The mechanism of plasmid transfer and the conditions necessary for transfer of chromosomal genes are therefore discussed in Chapter 28 , Bacterial Genetics.

Some plasmids can transfer themselves between bacterial cells and a few can also transfer chromosomal genes.

2.1 Plasmid Families and Incompatibility

Two different plasmids that belong to the same family cannot co-exist in the same cell. This is known as incompatibility . Plasmids were originally classified by incompatibility and so plasmid families are often known as incompatibility groups and are designated by letters of the alphabet (F, P, I, X, etc.). Plasmids of the same incompatibility group have very similar DNA sequences in their replication and partition genes, although the other genes they carry may be very different. It is quite possible to have two or more plasmids in the same cell as long as they belong to different families. So a P-type plasmid will happily share the same cell with a plasmid of the F-family ( Fig. 23.03 ).

Figure 23.03 . Plasmid Incompatibility

Plasmids with different origins of replication and different replication genes are able to inhabit the same bacterial cell and are considered compatible (left). During cell division, both types of plasmid replicate therefore, each daughter cell will inherit both plasmids, just like the mother cell. On the other hand, if two plasmids have identical origins and replication genes they are incompatible and will not be replicated during cell division (right). Instead, the two plasmids are partitioned into different daughter cells.

Plasmids are classified into families whose members share very similar replication genes.

2.2 Occasional Plasmids Are Linear or Made of RNA

Although most plasmids are circular molecules of DNA there are occasional exceptions. Linear plasmids of double-stranded DNA have been found in a variety of bacteria and in fungi and higher plants. The best-characterized linear plasmids are found in those few bacteria such as Borrelia and Streptomyces that also contain linear chromosomes. Linear DNA replicons in bacteria are not protected by telomeres like the linear chromosomes of eukaryotes. Instead, a variety of individual adaptations protect the ends from endonucleases.

Linear plasmids have special structures to protect the ends of the DNA.

In Borrelia, there are not actually any free DNA ends. Instead, hairpin sequences of single-stranded DNA form loops at the ends of both linear plasmids and chromosomes ( Fig. 23.04A ). Some animal viruses, such as the iridovirus that causes African swine fever, have similar structures. Different species of Borrelia cause Lyme’s disease and relapsing fever. Their linear plasmids appear to encode both hemolysins that damage blood cells and surface proteins that protect the bacteria from the host immune system. Thus, as is true of many other infectious bacteria, the virulence factors of Borrelia are also largely plasmid-borne.

Figure 23.04 . End Structures of Linear Plasmids

(A) Linear plasmids of Borrelia form hairpin loops at the ends. (B) Linear plasmids of Streptomyces are coated with proteins that protect the DNA ends. If linear plasmids had exposed double-stranded ends, this would trigger recombination, repair, or degradation systems.

The linear plasmids of Streptomyces are indeed genuine linear DNA molecules with free ends. They have inverted repeats at the ends of the DNA that are held together by proteins. In addition, special protective proteins are covalently attached to the 5′ ends of the DNA. The net result is a tennis racket structure ( Fig. 23.04B ). The DNA of adenovirus, most linear eukaryotic plasmids, and some bacterial viruses show similar structures.

Linear plasmids are also found among eukaryotes. The fungus Flammulina velutipes, commonly known as the enoki mushroom, has two very small linear plasmids within its mitochondria. Several higher plants are also known that have linear plasmids in their mitochondria. The dairy yeast, Kluyveromyces lactis, has a linear plasmid that normally replicates in the cytoplasm. However, on occasion the plasmid relocates to the nucleus where it replicates as a circle. Circularization is due to site specific recombination involving the inverted repeats at the ends of the linear form of the plasmid. The physiological role of these plasmids is obscure.

RNA plasmids are rare and most are poorly characterized. Examples are known from plants, fungi, and even animals. Some strains of the yeast Saccharomyces cerevisiae contain linear RNA plasmids. Similar RNA plasmids are found in the mitochondria of some varieties of maize plants. RNA plasmids are found as both single-stranded and double-stranded forms and replicate in a manner similar to certain RNA viruses. The RNA plasmid encodes RNA-dependent RNA polymerase that directs its own synthesis. Unlike RNA viruses, RNA plasmids do not contain genes for coat proteins. Sequence comparisons suggest that most RNA plasmids are merely defective versions of RNA viruses that have taken up permanent residence after losing the ability to move from cell to cell as virus particles.

How to spot plasmid on paper for shipping? - (Feb/09/2006 )

I need to send out a plasmid (to one of our lab's collaborators) by blotting on filter paper. Can somebody tell me the exact procedure for doing it: what's the amount of DNA to be spotted, which paper to use, how long to let it sit, etc. Thanks.

You can just spot your plasmid on a piece of filter paper. Before spotting, you can draw a circle on the paper using a pencil so that the recepitent will know where it is. There is no real limit on the amount of plasmid you can spot, it all depends on the concentration of your DNA.

So do you soak the piece of filter paper in comp cells and transform or what?

I've heard of this technique before but I have not tried it.

So do you soak the piece of filter paper in comp cells and transform or what?

I've heard of this technique before but I have not tried it.

you soak the piece of filter paper in TE (I take a sterile tip and smush around a little) then take say 5-10ul and transform.

hey, Matt, if you are looking for some technical references, this is commercially available as a product called Clonesaver

Experimental Procedure

Digest and purify vector:

While waiting for your oligos to arrive, conduct a restriction digest of 1μg of vector with EcoRI and SalI

Run an agarose gel and cut out the band containing your vector DNA

Gel purify your DNA away from the agarose using a commercially available kit or standard protocol.

Anneal oligos:

The oligos should be resuspended in annealing buffer (10mM Tris, pH7.5-8.0, 50mM NaCL, 1mM EDTA) and mixed in equimolar concentrations. We recommend mixing 2μg each in a total volume of 50μL - add additional annealing buffer if necessary to get to 50μL. Efficient annealing can be achieved by one of two methods:

Place the mixed oligos in a 1.5mL microfuge tube.

Place tube in 90-95°C hot block and leave for 3-5 minutes.

Remove the hot block from the heat source (turn off or move block to bench top) allowing for slow cooling to room temperature (

Place mixed oligos in a PCR tube.

Place tube in a thermocycler programmed to start at 95°C for 2 minutes.

Then, gradually cool to 25°C over 45 minutes.


Dilute 5μL of annealed oligos with 45μL nuclease-free water and quantify the concentration (should be about 8ng/μl).

Mix the annealed oligos with cut vector in molar ratios (vector:insert) between 4:3 and 1:6 in a standard ligation reaction (ex. to ligate an annealed oligo insert of 50bp in length into a 5kb vector, mix 100ng of the vector with 0.75-6 ng of annealed oligos).

Transform 2-3μL into your favorite competent bacteria and plate.

Be sure to pick multiple colonies for mini-prepping and verify insert by sequencing.

Types of control plasmids

Part of planning your experiment includes determining what factors need to be controlled for in order to eliminate any alternative interpretation of the results. Typically, plasmid-based experiments employ transfection, negative, positive, and replicate controls.

Transfection controls: empty vector and internal control

A transfection control measures transfection efficiency and enables observation of any effects of the transfection itself (i.e., the vector used in transfection, transfection reagent, or transfection process) may have on the target cells. One transfection control is an empty vector control specifically, the plasmid without the independent variable. Referring back to the experiment associated with Figure 1, the independent variable is the shRNA. Therefore the empty control vector would be Plasmid A sans shRNA, or backbone Y alone. The empty vector control allows you to examine if the transfection reagents or the transfection process itself has any cytotoxic effects on the target cells.

Another type of transfection control is an internal control vector, which measures transfection efficiency. An internal control may be a plasmid that constitutively expresses a reporter protein (e.g., GFP or luciferase) that is either co-transfected with the test plasmid or transfected into a separate well of your cells. Regardless, the amount of reporter protein activity correlates to both the amount of DNA transfected into the cells and the ability of the cells to express the protein.

In analysis of the result in Figure 1, an internal control, such as the GFP-expressing Plasmid B, could demonstrate whether the cells were transfected successfully and expressing the protein. For example, fluorescence microscopy images resulting from our experiment that includes the aforementioned internal control and is consistent with the result in Figure 1 could look like this:

Figure 2: Expression of Plasmid B (as internal control)

This result indicates that the transfection was not successful due to the absence of GFP fluorescence from the living, viable cells. This result presents an opportunity for troubleshooting and optimization of the transfection reagents and process. It is important to note that optimal experimental conditions, including how much plasmid DNA to use for any individual or co- transfection, should be determined empirically.

Negative controls: untreated cells, empty vector control, and non-targeting control

Negative control conditions and plasmids should produce a null effect (i.e., no phenomenon is observed). In any plasmid-based experiment, untreated cells should be included as these provide the baseline/standard against which other samples can be compared.

The Empty Vector Control (mentioned above) could also serve as an important negative control. In this context, the empty vector control shows any effect of the vector/backbone itself on gene expression in your target cells.

In experiments employing gene targeting or genome editing technologies, such as RNAi or CRISPR, non-targeting controls may be appropriate as they allow you to assess the specificity of your observed result. Non-targeting controls are negative controls that produce a similar product, but do not target an endogenous gene in your experimental cells. For example, in the experiment above, Plasmid A contains an shRNA that targets human Gene X and human cells are being transfected. A proper non-targeting control in this experiment could be an shRNA—in the same backbone as Plasmid A—that does not target any mammalian gene. This non-targeting control is critical to the correct interpretation of the results because it provides an important reference point when analyzing the specificity of the shRNA targeting human Gene X. The non-targeting control also assesses any effects of the general introduction of shRNA into your target cells.

Positive controls

Positive control plasmids should produce the expected phenomenon. One example of a positive control, the internal control vector, was described earlier. Once you are sure your conditions are conducive to the successful delivery of plasmid DNA into the cells, the internal control vector then serves as a positive control for transfection because it produces the expected effect, which is green fluorescent cells (Figure 3).

Figure 3: Expression of Plasmid B (as positive control)

Other positive controls are specific to the experiment and should be designed accordingly . If you are trying activate a gene, you should design a control that shows maximal activation. Likewise if you are trying to repress a gene, your control might be a system where expression of that gene is knocked out completely. These controls may or may not be plasmid-based depending on the experimental needs. Using our experiment as an example, the expected result of the shRNA targeting human Gene X in Plasmid A is the decreased expression of Gene X. Ergo the positive control(s) should decrease the expression of Gene X.

Since the key feature of a control plasmid is to minimize the effects of the non-independent variables within an experiment, both the design and selection of the positive control plasmids should be highly specific to the experiment and the interrogation of the independent variable.

How Does Replication Start and Stop?

In addition to genes, plasmids often include transcription promoters and terminators derived from E.coli phages. Promoters from phages SP6 and T7 are often used for in vitro RNA amplification. They require phage polymerases and are therefore inactive in vivo.

Below is a map of pTLNX , a Xenopus oocyte expression vector (Figure 2). In addition to the familiar pBR322 origin and antibiotic resistance genes AmpR and CmR (chloramphenicol resistance), there are also SP6 and lacUV promoters present. Downstream from SP6 promoter, the rrnBT2 terminator allows efficient termination of genes cloned into the multiple cloning site 2 (Figure 2).

The pTLNX vector also has a gene for plasmid selection (ccdB), along with virus SV40 nuclear localization signal and Xenopus globine 3’ UTR, that allows for high expression levels of cloned genes.

Many bacteria contain extra-chromosomal DNA elements called plasmids. These are usually small (a few 1000 bp), circular, double stranded molecules that replicate independently of the chromosome and can be present in high copy numbers within a cell. In the wild, plasmids can be transferred between individuals during bacterial mating and are sometimes even transferred between different species. Plasmids are particularly important in medicine because they often carry genes for pathogenicity and drug-resistance. In the lab, plasmids can be inserted into bacteria in a process called transformation.

To insert a DNA fragment into a plasmid, both the fragment and the circular plasmid are cut using a restriction enzyme that produces compatible ends (Figure (PageIndex<1>)). Given the large number of restriction enzymes that are currently available, it is usually not too difficult to find an enzyme for which corresponding recognition sequences are present in both the plasmid and the DNA fragment, particularly because most plasmid vectors used in molecular biology have been engineered to contain recognition sites for a large number of restriction endonucleases.

Figure (PageIndex<1>): Cloning of a DNA fragment (red) into a plasmid vector. The vector already contains a selectable marker gene (blue) such as an antibiotic resistance gene. (Original-Deyholos-CC:AN)

After restriction digestion, the desired fragments may be further purified or selected before they are mixed together with ligase to join them together. Following a short incubation, the newly ligated plasmids, containing the gene of interest are transformed into E. coli. Transformation is accomplished by mixing the ligated DNA with E. coli cells that have been specially prepared (i.e. made competent) to uptake DNA. Competent cells can be made by exposure to compounds such as CaCl2 or to electrical fields (electroporation). Because only a small fraction of cells that are mixed with DNA will actually be transformed, a selectable marker, such as a gene for antibiotic resistance, is usually also present on the plasmid. After transformation (combining DNA with competent cells), bacteria are spread on a bacterial agar plate containing an appropriate antibiotic so that only those cells that have actually incorporated the plasmid will be able to grow and form colonies. This can then be picked and used for further study.

Molecular biologists use plasmids as vectors to contain, amplify, transfer, and sometimes express genes of interest that are present in the cloned DNA. Often, the first step in a molecular biology experiment is to clone (i.e. copy) a gene into a plasmid, then transform this recombinant plasmid back into bacteria so that essentially unlimited copies of the gene (and the plasmid that carries it) can be made as the bacteria reproduce. This is a practical necessity for further manipulations of the DNA, since most techniques of molecular biology are not sensitive enough to work with just a single molecule at a time. Many molecular cloning and recombination experiments are therefore iterative processes in which:

  1. a DNA fragment (usually isolated by PCR and/or restriction digestion) is cloned into a plasmid cut with a compatible restriction enzyme
  2. the recombinant plasmid is transformed into bacteria
  3. the bacteria are allowed to multiply, usually in liquid culture
  4. a large quantity of the recombinant plasmid DNA is isolated from the bacterial culture
  5. further manipulations (such as site directed mutagenesis or the introduction of another piece of DNA) are conducted on the recombinant plasmid
  6. the modified plasmid is again transformed into bacteria, prior to further manipulations, or for expression

An Application of Molecular Cloning: Production of Recombinant Insulin

Purified insulin protein is critical to the treatment of diabetes. Prior to

1980, insulin for clinical use was isolated from human cadavers or from slaughtered animals such as pigs. Human-derived insulin generally had better pharmacological properties, but was in limited supply and carried risks of disease transmission. By cloning the human insulin gene and expressing it in E. coli, large quantities of insulin identical to the human hormone could be produced in fermenters, safely and efficiently. Production of recombinant insulin also allows specialized variants of the protein to be produced: for example, by changing a few amino acids, longer-acting forms of the hormone can be made. The active insulin hormone contains two peptide fragments of 21 and 30 amino acids, respectively. Today, essentially all insulin is produced from recombinant sources (Figure (PageIndex<2>)), i.e. human genes and their derivatives expressed in bacteria or yeast.

Figure (PageIndex<2>): A vial of insulin. Note that the label lists the origin as &ldquorDNA&rdquo, which stands for recombinant DNA.(Flickr-DeathByBokeh-CC:AN)

A plasmid is a circular double-stranded DNA molecule usually found in bacteria that is capable of replicating independently of the cell’s chromosomal DNA. Usually plasmids allow the bacteria to develop some advantage such as antibiotic resistance. A bacterial cell can have more than one plasmid, will express the genes on those plasmids and will replicate each plasmid when it divides. In this way each daughter cell will get a copy of the plasmid. Plasmids are often used for cloning purposes since genes of interest (fragments of DNA) can be inserted into plasmids and introduced into the bacteria by bacterial transformation. In bacterial transformation, competent bacteria where the cell wall has been made permeable to genetic material can take up a foreign plasmid. The bacteria that have taken up the plasmid can be selected for usually by growing the bacteria in a certain antibiotic to which the plasmid DNA allows resistance. In this way, as the bacteria divide the plasmid and thus the genes on them will be amplified.

These are enzymes that cut DNA at specific recognition sites that are usually 4 to 8 base pairs in length. The sites are usually also palindromic, meaning they read the same forwards and backwards. The restriction enzymes will also produce “sticky” or “blunt” ends. These ends can be used to insert the gene of interest into a plasmid by ligation.

Ex. Sticky end where bases are left unpaired after a cut

Ex. Blunt end where all bases are paired after a cut


The CGSC Database of E. coli genetic information includes genotypes and reference information for the strains in the CGSC collection, the names, synonyms, properties, and map position for genes, gene product information, and information on specific mutations and references to primary literature. The public version of the database includes this information and can be queried directly via this CGSC DB WebServer. For help, use the help links located above and on each query form, or contact us directly.

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