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New England Biolabs offers a wide variety of exonucleases with a range of nucleotide structure specificity. Exonucleases can be active on ssDNA and/or dsDNA, initiate from the 5´ end and/or the 3´ end of polynucleotides, and can also act on RNA. Exonucleases have many applications in molecular biology, including removal of PCR primers, cleanup of plasmid DNA and production of ssDNA from dsDNA. In this article, we explore the activity of commercially available exonucleases on oligonucleotides that have chemical modifications added during phosphoramidite synthesis, including phosphorothioate diester bonds, 2´-modified riboses, modified bases, and 5´ and 3´ end modifications. We discuss how modifications can be used to selectively protect some polynucleotides from digestion in vitro, and which modifications will be cleaved like natural DNA. This information can be helpful for designing primers that are stable to exonucleases, protecting specific strands of DNA, and preparing oligonucleotides with modifications that will be resistant to rapid cleavage by common exonuclease activities.
A variety of DNA exonucleases have been characterized from many different organisms; in vivo, these enzymes play critical roles in polynucleotide repair, recycling, error correction, and protection from exogenous DNA (6-8). In vitro, exonucleases are used in many applications where it is desirable to remove certain nucleic acids. For example, Exonuclease V (RecBCD) (Exo V, NEB #M0345) is often used to remove contaminating linear ssDNA and dsDNA from plasmid preparations (4,9); T7 Exonuclease (T7 Exo, NEB #M0263) can be used to generate 3´ overhangs in DNA (4, 10, 11); Exonuclease I (Exo I, NEB #M0293), Thermolabile Exonuclease I (NEB #M0568) or Exonuclease VII (Exo VII, NEB #M0379) can be used to eliminate ssDNA PCR primers, leaving double-stranded products undigested (12, 13), and Lambda Exonuclease (Lambda Exo, NEB #M0262) can be used to convert dsDNA to ssDNA for a variety of applications (14-16). More information on common applications of exonucleases available from NEB can be found in our selection chart, Common Applications of Exonucleases and Non-specific Endonucleases, at go.neb.com/ExosEndos.
Figure 1:
Examples of exonuclease directionality
Figure 2:
Examples of common nucleotide modifications and their effect on exonuclease activity
Figure 3:
Chirality of phosphorothioate bonds
Figure 4:
Designing oligonucleotides with nuclease-resistant modifications
(A) End fluorescein (FAM) labeled-DNA is rapidly degraded by exonucleases. (B) pt bonds between nucleotides prevent the DNA strand from being degraded, but the end label can still be cleaved. (C) An internal FAMdT surrounded by pt bonds will prevent the exonuclease from removing the label.
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Pictured are double stranded exonucleases with a 3´ to 5´ polarity (top), a 5´ to 3´ polarity (middle), and a bidirectional nuclease (bottom).
Modified phosphodiesters (pt bonds) block exonuclease activity
5´ and 3´ end modification does not block exonuclease activity
Modified bases do not inhibit exonuclease activity
Sugar modifications inhibit most exonuclease activity
Phosphorothioates can block many, but not all, exonucleases. To block exonuclease cleavage, the pt bonds must be placed at the end(s) where the enzyme initiates, e.g., the 5´ end for Lambda Exo and the 3´ end for Exo III. It is important to note that a single pt bond is insufficient to fully protect an oligonucleotide from exonuclease digestion. When the pt bond is installed via an oxidation step during phosphoramidite synthesis, a nearly equal amount of each isomer (SP and RP) is formed at each pt linkage (20). Since most enzymes can cleave one of these isomers, a single chemically installed pt will protect only half the molecules from digestion by a given exonuclease. Thus, it is typically recommended that 3–6 pt bonds be used to block exonuclease digestion, to prevent this read-through. One might expect that because each bond is a 50:50 mixture of isomers, when presented with 5 consecutive isomers, a given enzyme could cleave the first bond on half the molecules, then half of the molecules that had the first bond hydrolyzed would have the second hydrolyzed, and so on, such that there would be a range of partially degraded products. In practice, it has been reported (and confirmed by recent results at NEB) that five consecutive pt bonds completely block all exonuclease activity at all pt bond positions (16). The exact reasons for this are not currently known, but it is likely that exonucleases engage multiple bases at once, and the net effect of the isomeric mixture somehow prevents the active site from properly organizing around bonds that are the normally cleavable pt isomer. There are several commonly used exonucleases that are not blocked even by 5 consecutive pt bonds; for example, Exo V, Exo VII and T5 Exonuclease (T5 Exo, NEB #M0363) all can cleave, leaving short oligos instead of cutting at every bond in a series, and thus can digest DNA by skipping over termini blocked by multiple pt bonds and cleaving at the first phosphodiester (5, 26). Importantly, any enzyme with endonuclease activity, like DNase I, will simply ignore the ends and degrade the polynucleotides from the inside out (unless every phosphodiester bond is replaced by a phosphorothioate). Keeping these important exceptions in mind, phosphorothioate bonds remain the most generally applicable (and relatively inexpensive) way to protect oligonucleotides from digestion by exonucleases. For a complete list of DNA exonucleases and their interaction with pt bonds, view our selection chart, Activity of Exonucleases and Non-Specific Endonucleases, at go.neb.com/ExosEndos.
References 1. Lehman, I. R., and Nussbaum, A. L. (1964) J. Biol. Chem. 239, 2628-2636. 2. Nichols, N. M. (2011) Curr. Protoc. Mol. Biol. Chapter 3, Unit3.12. 3. Richardson, C. C., Lehman, I. R., and Kornberg, A. (1964) J. Biol. Chem. 239, 251-258. 4. McReynolds, L. A., and Nichols, N. M. (2011) Curr. Protoc. Mol. Biol. Chapter 3, Unit 3.11. 5. Lovett, S. T. (2011) EcoSal Plus 4. 6. Yang, W. (2011) Q. Rev. Biophys. 44, 1-93. 7. Bebenek, A., and Ziuzia-Graczyk, I. (2018) Curr. Genet. 64, 985-996. 8. Tsutakawa, S. E., Lafrance-Vanasse, J., and Tainer, J. A. (2014) DNA Repair (Amst). 19, 95-107. 9. Karu, A. E., MacKay, V., Goldmark, P. J., and Linn, S. (1973) J. Biol. Chem. 248, 4874-4884. 10. Kerr, C., and Sadowski, P. D. (1972) J. Biol. Chem. 247, 305-310. 11. Straus, N. A., and Zagursky, R. J. (1991) Biotechniques. 10, 376-384. 12. Li, H. H., Cui, X. F., and Arnheim, N. (1991) Nucleic. Acids. Res. 19, 3139-3141. 13. Enzymatic PCR Cleanup using Exonuclease I and Shrimp Alkaline Phosphatase. New England Biolabs, Ipswich, MA. 14. Civit, L., Fragoso, A., and O’Sullivan, C. K. (2012) Anal. Biochem. 431, 132-138. 15. Murgha, Y. E., Rouillard, J. M., and Gulari, E. (2014) PLoS One 9, e94752. 16. Nikiforov, T. T., Rendle, R. B., Kotewicz, M. L., and Rogers, Y. H. (1994) PCR Methods Appl. 3, 285-291. 17. Skerra, A. (1992) Nucleic Acids Res. 20, 3551-3554. 18. Evers, M. M., Toonen, L. J., and van Roon-Mom, W. M. (2015) Adv. Drug Deliv. Rev. 87, 90-103. 19. Lundin, K. E., Gissberg, O., and Smith, C. I. (2015) Hum. Gene Ther. 26, 475-485. 20. Eckstein, F. (1985) Annu. Rev. Biochem. 54, 367-402. 21. Spitzer, S., and Eckstein, F. (1988) Nucleic Acids Res. 16, 11691-11704. 22. Eckstein, F., and Gish, G. (1989) Trends Biochem Sci 14, 97-100. 23. Eckstein, F. (2014) Nucleic Acid Ther. 24, 374-387. 24. Putney, S. D., Benkovic, S. J., and Schimmel, P. R. (1981) Proc. Natl. Acad. Sci. U S A, 78, 7350-7354. 25. Yang, Z., Sismour, A. M., and Benner, S. A. (2007) Nucleic Acids Res. 35, 3118-3127. 26. Sayers, J. R., and Eckstein, F. (1990) J. Biol. Chem. 265, 18311-18317. 27. Kratschmer, C., and Levy, M. (2017) Nucleic Acid Ther. 27, 335-344. 28. Piccirilli, J. A., Krauch, T., Moroney, S. E., and Benner, S. A. (1990) Nature, 343, 33-37.
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Base modifications None of the exonucleases available from NEB were significantly inhibited by modified bases under the conditions we tested. Modifications tested included 5-methyl-substituted dT (e.g., Fluorescein dT), deoxyuridine, the Tm-enhancing “super T,” and the non-natural base pair isoG:isoC (Figure 2) (28). All modified substrates were digested completely by all the exonucleases tested. Some modifications showed weak blockage, pausing at the modification site before completely degrading the substrate. For several exonucleases tested, modified dT bases with large modifications off the 5-methyl position (Figure 2) showed a buildup of partially-digested intermediates, apparently stalling just before the modification; in no case did this resistance approach the inhibition seen for 2´ MOE sugars or pt linkages. Conclusion We have evaluated a variety of chemical modifications for their inhibition of exonuclease activity at the 5´ and 3´ ends of oligonucleotides. Broadly, the phosphorothioate modification, one of the more well-known used modifications to block nuclease cleavage, remains the most effective choice to protect oligonucleotides from degradation. However, one must be careful to use multiple pt bonds, place them at the correct end of the oligonucleotide to match the polarity of the exonucleases used, and be aware that several exonucleases can read-through or bypass terminal pt bonds; your choice of nucleases is as important as the modifications used. Aside from pt bonds, MOE nucleotides are the next best choice for providing nuclease resistance in vitro, with similar caveats to pt bonds. The vast majority of end modifications, including affinity tags and fluorophores, as well as internal non-standard bases, provide little, if any, nuclease resistance, and will be cleaved completely in vitro.
2´-modified nucleosides Generally, DNA exonucleases do not digest RNA portions of oligonucleotides, though RNA is itself susceptible to RNases and nonspecific hydrolysis. We have further found that hybridizing RNA to DNA strands does not block the activity of dsDNA exonucleases on the DNA strand. Hybridization of ssDNA to RNA will block the activity of ssDNA exonucleases as effectively as hybridization to dsDNA. Additionally, certain 2´-O-modified riboses, are both stable to spontaneous hydrolysis and offer strong resistance to exonuclease activity (27). 2´-O-methyl and 2´-O-methoxyethyl (MOE) nucleosides, which contain bulky substituents off the sugar ring, have been shown to grant strong resistance to nucleases and additionally increase the strength of annealing to complementary DNA and RNA. These features have found utility in antisense nuclease strategies, to make oligonucleotides that are both resistant to degradation and able to bind tightly to target RNAs. These sugar modifications also work in vitro to block exonuclease activity quite strongly. Our studies have found that, while a single terminal MOE nucleoside only weakly inhibits exonuclease activity, three successive MOE modifications provide enhanced resistance to many exonucleases, including Exo I, Exo III, Lambda Exo, RecJf (NEB #M0264) and polymerase exonucleases, such as that of DNA Polymerase I, Large (Klenow) Fragment (NEB #M0210). Similar to pt bonds, several exonucleases can digest through these regions, notably T5 Exo, T7 Exo, Exo V, Exo VII and Exo VIII. Overall, exonuclease inhibition by MOE is quite strong, but pt bonds are more effective and are typically cheaper to prepare and incorporate. However, if for some reason the pt chemistry is not desired, 2´-O-modified ribose moieties are a viable alternative.
The ability of nucleases to hydrolyze phosphodiester bonds in nucleic acids is among the earliest nucleic acid enzyme activities to be characterized (1-6). Endonucleases cleave internal phosphodiester bonds, while exonucleases, the focus of this article, must begin at the 5´ or 3´ end of a nucleic acid strand and cleave the bonds sequentially (Figure 1). Exonucleases may be DNA or RNA specific, and can act on single-stranded or double-stranded nucleic acids, or both. Double-strand specific exonucleases may initiate at blunt ends, nicks, or short single-stranded 5´ or 3´ overhangs, though most exonucleases are active on a subset of these structures. For a summary of the substrate specificity of exonucleases available from NEB, view our newly-updated selection chart, Properties of Exonucleases and Non-specific Endonucleases, at go.neb.com/ExosEndos.
Phosphorothioate linkages A phosphorothioate (pt) bond is a phosphodiester linkage where one of the two non-bridging oxygens has been replaced by a sulfur (Figure 2). This modification has been used for decades to inhibit nuclease phosphodiesterase and phosphoryl transferase activities, as well as for gaining mechanistic insights into these enzymes (20, 23). Chemically, the substitution of oxygen with sulfur does not dramatically change the reactivity of the bond, and pt-containing polynucleotides can still function in many enzymatic reactions. In a typical phosphodiester bond, the two non-bridging oxygens are chemically equivalent. When one of these oxygens is replaced by sulfur, however, the phosphorus is now connected to four distinct groups, rendering it a chiral center with two possible configurations referred to as “SP” and “RP” (Figure 3). It is this key feature that confers resistance for the majority of nuclease enzymes studied; one configuration will react at rates similar to a phosphodiester, while the other is significantly inhibitory or completely unreactive. Isomer reactivity varies from enzyme to enzyme, and different pt isomers can inhibit enzymes that catalyze the same reaction (e.g., phosphoryl transfer). For example, DNA Polymerase I (DNA Pol I, NEB #M0209) can incorporate deoxynucleotide triphosphates with a pt ester at the α phosphate (dNTPαS), allowing formation of pt-bonded polynucleotides. However, it can only react with SP configured dNTPαS molecules, and does so with inversion of the stereocenter to form exclusively RP-configured pt bonds in the product. Conversely, the 3´→ 5´exo activity of this polymerase cleaves RP, but not SP configured bonds (20). Alternatively, the 3´→ 5´ exo activity of E. coli Exonuclease III (Exo III, NEB #M0206) cleaves SP, but not RP configured pt bonds (24). Therefore, DNA created from the incorporation of dNTP S by DNA Pol I is highly resistant to exonuclease cleavage by Exo III (25).
A variety of DNA exonucleases have been characterized from many different organisms; in vivo, these enzymes play critical roles in polynucleotide repair, recycling, error correction, and protection from exogenous DNA (6-8). In vitro, exonucleases are used in many applications where it is desirable to remove certain nucleic acids. For example, Exonuclease V (RecBCD) (Exo V, NEB #M0345) is often used to remove contaminating linear ssDNA and dsDNA from plasmid preparations (4,9); T7 Exonuclease (T7 Exo, NEB #M0263) can be used to generate 3´ overhangs in DNA (4, 10, 11); Exonuclease I (Exo I, NEB #M0293), Thermolabile Exonuclease I (NEB #M0568) or Exonuclease VII (Exo VII, NEB #M0379) can be used to eliminate ssDNA PCR primers, leaving double-stranded products undigested (12, 13), and Lambda Exonuclease (Lambda Exo, NEB #M0262) can be used to convert dsDNA to ssDNA for a variety of applications (14-16). More information on common applications of exonucleases available from NEB can be found in our selection chart, Common Applications of Exonucleases and Non-specific Endonucleases, at go.neb.com/ExosEndos, What about cases where you only want to degrade some of the ssDNA in a reaction? Or, when you want to make ssDNA from a dsDNA substrate, but which strand is degraded matters greatly? What about cases where the ends of your nucleic acids are modified—will exonucleases still digest the substrate, or cleave the modification? Several methods depend on selective protection of polynucleotides, such as protection of primers from degradation by polymerase exonuclease domains (17), selective protection of one strand of a DNA duplex for the production of ssDNA (14-16), and the protection of polynucleotides from degradation by serum nucleases, as in the case of RNA interference drugs (18, 19). In each of these cases, it is critical to understand the influence of modifications on exonuclease activity—which modifications inhibit nucleotide cleavage and which do not. Recently, researchers at NEB have worked to characterize the interaction between exonucleases and modified polynucleotides, as part of a broader effort to gain deeper insight into the sequence and structural determinants of nuclease activity and specificity. In an effort to catalog the modifications that inhibit exonuclease digestion, we treated polynucleotides containing a range of modifications (including non-standard bases, sugars and backbone chemistries) with exonucleases under the recommended in vitro reaction conditions. This article will summarize data from the literature, as well as the key results from NEB’s work related to understanding the activity of exonucleases on chemically modified polynucleotides. We will focus on the most widely used—and most successful—method for blocking nuclease activity, the phosphorothioate bond (20-23), but will also discuss the use of other modifications to inhibit nuclease activity, as well as which modifications have little to no effect on exonuclease digestion. Phosphorothioate linkages A phosphorothioate (pt) bond is a phosphodiester linkage where one of the two non-bridging oxygens has been replaced by a sulfur (Figure 2). This modification has been used for decades to inhibit nuclease phosphodiesterase and phosphoryl transferase activities, as well as for gaining mechanistic insights into these enzymes (20, 23). Chemically, the substitution of oxygen with sulfur does not dramatically change the reactivity of the bond, and pt-containing polynucleotides can still function in many enzymatic reactions. In a typical phosphodiester bond, the two non-bridging oxygens are chemically equivalent. When one of these oxygens is replaced by sulfur, however, the phosphorus is now connected to four distinct groups, rendering it a chiral center with two possible configurations referred to as “SP” and “RP” (Figure 3). It is this key feature that confers resistance for the majority of nuclease enzymes studied; one configuration will react at rates similar to a phosphodiester, while the other is significantly inhibitory or completely unreactive. Isomer reactivity varies from enzyme to enzyme, and different pt isomers can inhibit enzymes that catalyze the same reaction (e.g., phosphoryl transfer). For example, DNA Polymerase I (DNA Pol I, NEB #M0209) can incorporate deoxynucleotide triphosphates with a pt ester at the α phosphate (dNTPαS), allowing formation of pt-bonded polynucleotides. However, it can only react with SP configured dNTPαS molecules, and does so with inversion of the stereocenter to form exclusively RP-configured pt bonds in the product. Conversely, the 3´→ 5´exo activity of this polymerase cleaves RP, but not SP configured bonds (20). Alternatively, the 3´→ 5´ exo activity of E. coli Exonuclease III (Exo III, NEB #M0206) cleaves SP, but not RP configured pt bonds (24). Therefore, DNA created from the incorporation of dNTP S by DNA Pol I is highly resistant to exonuclease cleavage by Exo III (25).
Other 5´/3´ end modifications Several other modifications, such as the inverted deoxythymidine bases and dideoxynucleotides (Figure 2) have been reported to suppress serum nuclease activity when appended to the end of synthetic oligonucleotides (27). Many other modifications may be attached through “linkers” at either the 5´ or 3´ end, including fluorescent tags, biotin or other affinity labels, or reactive groups for attachment to beads or surfaces. These linkers are typically connected to the 5´ or 3´ end via a phosphodiester, but what is the interaction of these modified ends with exonucleases? We have surveyed a range of these modifications in typical in vitro exonuclease assays. In general, while many provide modest inhibition as compared to a 5´-phosphate, all exonucleases tested could cleave all modifications connected through phosphodiester bonds. Interestingly, this poor inhibition held true for the inverted dT modifications, which have been reported to grant extra stability versus degradation by serum exonucleases for aptamers and other modified oligonucleotides. In our hands, 3´-inverted dT blocked only the relatively weak 3´→ 5´ exonuclease activity of DNA Polymerase I, Large (Klenow) Fragment (NEB #M0210) and Exonuclease T (Exo T, NEB #M0265), but did not block more active exonucleases such as in T7 DNA Polymerase (NEB #M0274), Exo I or Exo III. Similarly, 5´-inverted dT partially inhibited only Lambda Exo activity, which is known to require a 5´-phosphate for efficient initiation. Other 5´→ 3´ exonucleases were not significantly inhibited by this modification, showing complete digestion after a one-hour incubation under the recommended usage conditions. We do not recommend 5´/3´ end modification as a good strategy for producing nucleotides resistant to exonuclease degradation in vitro. Researchers should be aware that these modifications will be cleaved by the majority of exonucleases, potentially leading to the loss of fluorescent labels and affinity tags. If a modification stable to exonuclease activity is needed, a better strategy is to use internal labels connected to the 5-methyl position of dT (e.g., Fluorescein dT, Figure 2). If these modified dT bases are used near the end of an oligo, they can be protected with surrounding pt bonds (Figure 4). The linkage to the base is not susceptible to enzymatic cleavage, and the pt bonds will protect the backbone from digestion, as described above. Base modifications None of the exonucleases available from NEB were significantly inhibited by modified bases under the conditions we tested. Modifications tested included 5-methyl-substituted dT (e.g., Fluorescein dT), deoxyuridine, the Tm-enhancing “super T,” and the non-natural base pair isoG:isoC (Figure 2) (28). All modified substrates were digested completely by all the exonucleases tested. Some modifications showed weak blockage, pausing at the modification site before completely degrading the substrate. For several exonucleases tested, modified dT bases with large modifications off the 5-methyl position (Figure 2) showed a buildup of partially-digested intermediates, apparently stalling just before the modification; in no case did this resistance approach the inhibition seen for 2´ MOE sugars or pt linkages. Conclusion We have evaluated a variety of chemical modifications for their inhibition of exonuclease activity at the 5´ and 3´ ends of oligonucleotides. Broadly, the phosphorothioate modification, one of the more well-known used modifications to block nuclease cleavage, remains the most effective choice to protect oligonucleotides from degradation. However, one must be careful to use multiple pt bonds, place them at the correct end of the oligonucleotide to match the polarity of the exonucleases used, and be aware that several exonucleases can read-through or bypass terminal pt bonds; your choice of nucleases is as important as the modifications used. Aside from pt bonds, MOE nucleotides are the next best choice for providing nuclease resistance in vitro, with similar caveats to pt bonds. The vast majority of end modifications, including affinity tags and fluorophores, as well as internal non-standard bases, provide little, if any, nuclease resistance, and will be cleaved completely in vitro.
A variety of DNA exonucleases have been characterized from many different organisms; in vivo, these enzymes play critical roles in polynucleotide repair, recycling, error correction, and protection from exogenous DNA (6-8). In vitro, exonucleases are used in many applications where it is desirable to remove certain nucleic acids. For example, Exonuclease V (RecBCD) (Exo V, NEB #M0345) is often used to remove contaminating linear ssDNA and dsDNA from plasmid preparations (4,9); T7 Exonuclease (T7 Exo, NEB #M0263) can be used to generate 3´ overhangs in DNA (4, 10, 11); Exonuclease I (Exo I, NEB #M0293), Thermolabile Exonuclease I (NEB #M0568) or Exonuclease VII (Exo VII, NEB #M0379) can be used to eliminate ssDNA PCR primers, leaving double-stranded products undigested (12, 13), and Lambda Exonuclease (Lambda Exo, NEB #M0262) can be used to convert dsDNA to ssDNA for a variety of applications (14-16). More information on common applications of exonucleases available from NEB can be found in our selection chart, Common Applications of Exonucleases and Non-specific Endonucleases, at go.neb.com/ExosEndos, What about cases where you only want to degrade some of the ssDNA in a reaction? Or, when you want to make ssDNA from a dsDNA substrate, but which strand is degraded matters greatly? What about cases where the ends of your nucleic acids are modified—will exonucleases still digest the substrate, or cleave the modification? Several methods depend on selective protection of polynucleotides, such as protection of primers from degradation by polymerase exonuclease domains (17), selective protection of one strand of a DNA duplex for the production of ssDNA (14-16), and the protection of polynucleotides from degradation by serum nucleases, as in the case of RNA interference drugs (18, 19). In each of these cases, it is critical to understand the influence of modifications on exonuclease activity—which modifications inhibit nucleotide cleavage and which do not. Recently, researchers at NEB have worked to characterize the interaction between exonucleases and modified polynucleotides, as part of a broader effort to gain deeper insight into the sequence and structural determinants of nuclease activity and specificity. In an effort to catalog the modifications that inhibit exonuclease digestion, we treated polynucleotides containing a range of modifications (including non-standard bases, sugars and backbone chemistries) with exonucleases under the recommended in vitro reaction conditions. This article will summarize data from the literature, as well as the key results from NEB’s work related to understanding the activity of exonucleases on chemically modified polynucleotides. We will focus on the most widely used—and most successful—method for blocking nuclease activity, the phosphorothioate bond (20-23), but will also discuss the use of other modifications to inhibit nuclease activity, as well as which modifications have little to no effect on exonuclease digestion.
Other 5´/3´ end modifications Several other modifications, such as the inverted deoxythymidine bases and dideoxynucleotides (Figure 2) have been reported to suppress serum nuclease activity when appended to the end of synthetic oligonucleotides (27). Many other modifications may be attached through “linkers” at either the 5´ or 3´ end, including fluorescent tags, biotin or other affinity labels, or reactive groups for attachment to beads or surfaces. These linkers are typically connected to the 5´ or 3´ end via a phosphodiester, but what is the interaction of these modified ends with exonucleases? We have surveyed a range of these modifications in typical in vitro exonuclease assays. In general, while many provide modest inhibition as compared to a 5´-phosphate, all exonucleases tested could cleave all modifications connected through phosphodiester bonds. Interestingly, this poor inhibition held true for the inverted dT modifications, which have been reported to grant extra stability versus degradation by serum exonucleases for aptamers and other modified oligonucleotides. In our hands, 3´-inverted dT blocked only the relatively weak 3´→ 5´ exonuclease activity of DNA Polymerase I, Large (Klenow) Fragment (NEB #M0210) and Exonuclease T (Exo T, NEB #M0265), but did not block more active exonucleases such as in T7 DNA Polymerase (NEB #M0274), Exo I or Exo III. Similarly, 5´-inverted dT partially inhibited only Lambda Exo activity, which is known to require a 5´-phosphate for efficient initiation. Other 5´→ 3´ exonucleases were not significantly inhibited by this modification, showing complete digestion after a one-hour incubation under the recommended usage conditions. We do not recommend 5´/3´ end modification as a good strategy for producing nucleotides resistant to exonuclease degradation in vitro. Researchers should be aware that these modifications will be cleaved by the majority of exonucleases, potentially leading to the loss of fluorescent labels and affinity tags. If a modification stable to exonuclease activity is needed, a better strategy is to use internal labels connected to the 5-methyl position of dT (e.g., Fluorescein dT, Figure 2). If these modified dT bases are used near the end of an oligo, they can be protected with surrounding pt bonds (Figure 4). The linkage to the base is not susceptible to enzymatic cleavage, and the pt bonds will protect the backbone from digestion, as described above.
A variety of DNA exonucleases have been characterized from many different organisms; in vivo, these enzymes play critical roles in polynucleotide repair, recycling, error correction, and protection from exogenous DNA (6-8). In vitro, exonucleases are used in many applications where it is desirable to remove certain nucleic acids. For example, Exonuclease V (RecBCD) (Exo V, NEB #M0345) is often used to remove contaminating linear ssDNA and dsDNA from plasmid preparations (4,9); T7 Exonuclease (T7 Exo, NEB #M0263) can be used to generate 3´ overhangs in DNA (4, 10, 11); Exonuclease I (Exo I, NEB #M0293), Thermolabile Exonuclease I (NEB #M0568) or Exonuclease VII (Exo VII, NEB #M0379) can be used to eliminate ssDNA PCR primers, leaving double-stranded products undigested (12, 13), and Lambda Exonuclease (Lambda Exo, NEB #M0262) can be used to convert dsDNA to ssDNA for a variety of applications (14-16). More information on common applications of exonucleases available from NEB can be found in our selection chart, Common Applications of Exonucleases and Non-specific Endonucleases, at go.neb.com/ExosEndos, What about cases where you only want to degrade some of the ssDNA in a reaction? Or, when you want to make ssDNA from a dsDNA substrate, but which strand is degraded matters greatly? What about cases where the ends of your nucleic acids are modified—will exonucleases still digest the substrate, or cleave the modification? Several methods depend on selective protection of polynucleotides, such as protection of primers from degradation by polymerase exonuclease domains (17), selective protection of one strand of a DNA duplex for the production of ssDNA (14-16), and the protection of polynucleotides from degradation by serum nucleases, as in the case of RNA interference drugs (18, 19). In each of these cases, it is critical to understand the influence of modifications on exonuclease activity—which modifications inhibit nucleotide cleavage and which do not. Recently, researchers at NEB have worked to characterize the interaction between exonucleases and modified polynucleotides, as part of a broader effort to gain deeper insight into the sequence and structural determinants of nuclease activity and specificity. In an effort to catalog the modifications that inhibit exonuclease digestion, we treated polynucleotides containing a range of modifications (including non-standard bases, sugars and backbone chemistries) with exonucleases under the recommended in vitro reaction conditions. This article will summarize data from the literature, as well as the key results from NEB’s work related to understanding the activity of exonucleases on chemically modified polynucleotides. We will focus on the most widely used—and most successful—method for blocking nuclease activity, the phosphorothioate bond (20-23), but will also discuss the use of other modifications to inhibit nuclease activity, as well as which modifications have little to no effect on exonuclease digestion. Phosphorothioate linkages A phosphorothioate (pt) bond is a phosphodiester linkage where one of the two non-bridging oxygens has been replaced by a sulfur (Figure 2). This modification has been used for decades to inhibit nuclease phosphodiesterase and phosphoryl transferase activities, as well as for gaining mechanistic insights into these enzymes (20, 23). Chemically, the substitution of oxygen with sulfur does not dramatically change the reactivity of the bond, and pt-containing polynucleotides can still function in many enzymatic reactions. In a typical phosphodiester bond, the two non-bridging oxygens are chemically equivalent. When one of these oxygens is replaced by sulfur, however, the phosphorus is now connected to four distinct groups, rendering it a chiral center with two possible configurations referred to as “SP” and “RP” (Figure 3). It is this key feature that confers resistance for the majority of nuclease enzymes studied; one configuration will react at rates similar to a phosphodiester, while the other is significantly inhibitory or completely unreactive. Isomer reactivity varies from enzyme to enzyme, and different pt isomers can inhibit enzymes that catalyze the same reaction (e.g., phosphoryl transfer). For example, DNA Polymerase I (DNA Pol I, NEB #M0209) can incorporate deoxynucleotide triphosphates with a pt ester at the α phosphate (dNTPαS), allowing formation of pt-bonded polynucleotides. However, it can only react with SP configured dNTP S molecules, and does so with inversion of the stereocenter to form exclusively RP-configured pt bonds in the product. Conversely, the 3´→ 5´exo activity of this polymerase cleaves RP, but not SP configured bonds (20). Alternatively, the 3´→ 5´ exo activity of E. coli Exonuclease III (Exo III, NEB #M0206) cleaves SP, but not RP configured pt bonds (24). Therefore, DNA created from the incorporation of dNTP S by DNA Pol I is highly resistant to exonuclease cleavage by Exo III (25).
Exonucleases and endonucleases are used in many of today’s molecular biology workflows and applications. Did you know that NEB offers the largest supply of these important tools, and has a team of experts studying the function and optimization of these enzymes? We also offer several helpful tools to help you find the best enzyme to facilitate your work, including selection charts, recommended applications, usage guidelines and more.
Featured DNA Repair Enzymes and Structure-specific Endonucleases
Thermolabile USER II Enzyme
®
Thermolabile USER (Uracil-Specific Excision Reagent) II Enzyme generates a single nucleotide gap at the location of a uracil residue. It can be 100% inactivated at temperatures >65°C, streamlining workflows and enabling DNA to be used directly in downstream applications.
Antarctic Thermolabile Uracil DNA glycosylase (UDG) catalyzes the excision of a uracil base, forming an abasic (apyrimidinic) site while leaving the phosphodiester backbone intact. The lyase activity of Endonuclease III breaks the phosphodiester backbone at the 3´and 5´ sides of the abasic site. In addition to generating a different 3´-terminus than USER Enzyme (a 3´-phospho-α, β-unsaturated aldehyde versus the 3´phosphate left by USER Enzyme, NEB #M5505), Thermolabile USER II Enzyme (NEB #M5508) can also be completely heat inactivated after 10 minutes at 65°C.
Applications include:
• Directional RNA-seq • NEBNext® adaptor cleavage • USER cloning
Order
Thermostable FEN1
Thermostable Flap Endonuclease 1, FEN1, catalyzes the cleavage of 5´ DNA flaps from branched double stranded DNA substrates, creating a 5´ phosphate terminus. FEN1 products can be ligated by DNA ligase to create double-stranded DNA. In vivo, FEN1 is an essential component of the Okazaki fragment maturation pathway, and also plays a role in base excision repair.
Application:
• Base excision repair
• Determining genome targeting efficiency (access protocol via the protocol tab at www.neb.com/M0302)
T7 Endonuclease I
T7 Endonuclease I recognizes and cleaves non-perfectly matched DNA, cruciform DNA structures, Holliday structures or junctions, heteroduplex DNA and more slowly, nicked double-stranded DNA. The cleavage site is at the first, second or third phosphodiester bond that is 5´ to the mismatch. The protein is the product of T7 gene 3.
• Conversion of linear double-stranded DNA to single-stranded DNA via preferred activity on 5´-phosphorylated ends
Lambda Exonuclease
Lambda Exonuclease catalyzes the removal of nucleotides from linear or nicked double-stranded DNA in the 5´ to 3´ direction. This enzyme is highly processive. The preferred substrate is 5´-phosphorylated double-stranded DNA, although non-phosphorylated substrates are degraded at a greatly reduced rate.
• Removal of single-stranded primers in PCR reactions prior to DNA sequencing or SNP analysis • Removal of single-stranded primers for nested PCR reactions • Removal of linear single-stranded DNA, leaving behind double-stranded DNA
Thermolabile Exonuclease I
Thermolabile Exonuclease I catalyzes the removal of nucleotides from linear single-stranded DNA in the 3´ to 5´ direction. Unlike Exonuclease I (NEB #M0293), Thermolabile Exonuclease I can be heat inactivated at 80°C in one minute.
Featured Resources
Request your copy of our endonuclease magnet or exonuclease poster at:
www.neb.com/ExosEndosRequest
View the full list of exonucleases available at NEB
View the full list of endonucleases available at NEB
View extensive selection charts and activity information
View our webinar
Additional Information
View as pdf
Common Applications for Exonucleases and Endonucleases
Not sure which exonuclease or endonuclease to choose?
APPLICATION: Removal of 3´ overhangs RECOMMENDED ENZYME: Quick Blunting Kit NEB#: E1201S/L APPLICATION: 5´ overhang fill-in treatment RECOMMENDED ENZYME: Quick Blunting Kit NEB#: E1201S/L APPLICATION: Removal of ss primers for nested PCR reactions RECOMMENDED ENZYME: Thermolabile Exonuclease I NEB#: M0568S/L APPLICATION: Removal of primers post PCR prior to DNA sequencing or SNP detection RECOMMENDED ENZYME: Exonuclease I, Thermolabile Exonuclease I (1), Exonuclease VII (2) NEB#: M0293S/L NEB#: M0568S/L NEB#: M0379S/L APPLICATION: Mapping positions of introns in genomic DNA RECOMMENDED ENZYME: Exonuclease VII NEB#: M0379S/L APPLICATION: Removal of primers with or without 3´ or 5´ terminal phosphorothioate bonds RECOMMENDED ENZYME: Exonuclease VII NEB#: M0379S/L APPLICATION: Generating ssDNA from linear dsDNA: If 5´ 3´ polarity required RECOMMENDED ENZYME: Lambda Exonuclease (3) NEB#: M0262S/L If 3´ 5´ polarity required RECOMMENDED ENZYME: Exonuclease III (4) NEB#: M0206S/L APPLICATION: Preparation of nested deletions in double-stranded DNA RECOMMENDED ENZYME: Exonuclease III (E. coli) plus NEB#: M0206S/L RECOMMENDED ENZYME: Exonuclease VII NEB#: M0379S/L APPLICATION: Site-directed mutagenesis RECOMMENDED ENZYME: Exonuclease III (E. coli) (5) NEB#: M0206S/L RECOMMENDED ENZYME: T7 Exonuclease (6) NEB#: M0263S/L APPLICATION: Nick-site extension RECOMMENDED ENZYME: T7 Exonuclease NEB#: M0263S/L APPLICATION: Degradation of denatured DNA from alkaline-based plasmid purification methods for improving DNA cloning RECOMMENDED ENZYME: T5 Exonuclease NEB#: M0363S/L APPLICATION: Removal of chromosomal/linear DNA in plasmid preparations RECOMMENDED ENZYME: T5 Exonuclease (7) NEB#: M0363S/L RECOMMENDED ENZYME: Exonuclease V (RecBCD) (8) NEB#: M0345S/L APPLICATION: Removal of unligated products (linear dsDNA) from ligated circular double-stranded DNA RECOMMENDED ENZYME: T5 Exonuclease (9) NEB#: M0363S/L RECOMMENDED ENZYME: Exonuclease V (RecBCD) (10) NEB#: M0345S/L APPLICATION: Removal of residual gDNA after purification of low copy plasmid RECOMMENDED ENZYME: Exonuclease V (RecBCD) NEB#: M0345S/L APPLICATION: Removal of contaminating DNA from RNA samples RECOMMENDED ENZYME: DNase I NEB#: M0303S/L APPLICATION: Conversion of single-stranded DNA or RNA to 5´-mononucleotides RECOMMENDED ENZYME: Nuclease P1 NEB#: M0660S APPLICATION: Analysis of base composition, potential damage and modification of nucleic acids RECOMMENDED ENZYME: Nuclease P1 NEB#: M0660S APPLICATION: Progressive shortening of both ends of double-stranded DNA RECOMMENDED ENZYME: Nuclease BAL-31 NEB#: M0213S APPLICATION: Preparation of double-stranded DNA fragments with 5´-OH and 3´-phosphate RECOMMENDED ENZYME: Micrococcal Nuclease NEB#: M0247S APPLICATION: Degradation of nucleic acids (both DNA and RNA) in crude cell-free extracts RECOMMENDED ENZYME: Micrococcal Nuclease NEB#: M0247S APPLICATION: Preparation of rabbit reticulocyte RECOMMENDED ENZYME: Micrococcal Nuclease NEB#: M0247S APPLICATION: Chromatin Immunoprecipitation (ChIP) analysis RECOMMENDED ENZYME: Micrococcal Nuclease NEB#: M0247S
Notes: 1. Rapid heat inactivation versus Exonuclease I 2. For 3´ chemically modified primers 3. Strand targeted for removal requires one 5´ end with phosphate 4. Strand targeted for removal requires a 5´ overhang, a blunt end, or a 3´ overhang (with less than 4 bases) 5. Removes nicked-strand DNA from 3´ to 5´ 6. Removes nicked-strand DNA from 5´ to 3´ 7. Degrades linear ss + dsDNA, nicked DNA 8. Degrades linear ss + dsDNA: preferred as Exo V will save nicked plasmids resulting in higher yields especially for low-copy number plasmid prep 9. Only the unnicked form of ligated circular double-stranded DNA remains 10. Both nicked and unnicked form of ligated circular double-stranded DNA remains
Find the right enzyme for your application using the list below.
2019 Passion in Science Award winners pictured outside the New England Biolabs facility in Ipswich, MA.
This past May, New England Biolabs held its third Passion in Science Awards to recognize those within the scientific community who share the same core values as NEB and are working to solve many of today’s challenges. We were pleased to welcome 12 award winners to the NEB campus in Ipswich, Massachusetts to celebrate their dedication to science mentorship, humanitarianism, environmental stewardship and artistic creativity, and to learn how scientists can create unique opportunities to explore their passions.
Scientific Mentorship and Advocacy
Sarah McAnulty University of Connecticut, Willimantic, CT, USA
Nathan Schoepp Caltech, Pasadena, CA, USA
Bryan Welm University of Utah, Salt Lake City, UT, USA
“As I developed as a scientist I began to realize the power of the scientific process. I felt emboldened by my growing ability to seek answers to my own questions in a methodical way. I find the most meaning and value in helping others learn to answer their own questions in critical ways.”
Sarah Fankhauser Oxford College of Emory University, Oxford, GA, USA
“Through this work, I hope to inspire a solution that would liberate science in Africa, as well as create the platform that would permit scientists like myself to return home and do good science.”
Mahmoud Bukar Maina University of Sussex, England, UK
View the full event and inspiring presentations at www.neb.com/PassionInScience
Samantha Romanick University of Nevada Reno Reno, NV, USA
Daniel Heid University of Heidelberg, Heidelberg, Germany
Steven Farber Carnegie Institution for Science, Baltimore, MD, USA
Malali Gowda TransDisciplinary University, Bangaluru, India
William Ward Murta Theater Bielefeld, Bielefeld, Germany
Garfield Kwan University of California, San Diego, CA, USA
Kyle McClary University of Southern California, Los Angeles, CA, USA
Humanitarian Duty
Arts and Creativity
Environmental Stewardship
“For young children, seeing themselves represented in science is essential to encourage them to not only pursue science, but trust that scientists can be like them.” Skype a Scientist Sarah is the founder of Skype a Scientist, which connects non-scientists with scientists via personal video chats, enabling students and adults to learn about a scientist’s life, work and passions. Sarah has built a team of volunteer scientists and communicators who have served over 13,000 groups with live-stream or in-person presentations, as well as question and answer sessions. Skype a Scientist matches classrooms with scientists, and also offers “Skype a Scientist Live” sessions on YouTube Live, featuring high-profile scientists communicating their work and answering the public’s questions.
“The Xenoplex has often been the first step to a life as a scientist. Working in a laboratory is a perfect way to proof their skills and interests in an important work-orientation phase of their life, but also to learn a lot on how to organize scientific projects and present them to the public.” Xenoplex STEM Center Daniel is the founder of the Xenoplex STEM Center, which encourages young students who are interested in science and equips them with the experience and expertise to solve future scientific problems. This supra-regional learning center in Southern Germany pairs highly talented students with mentors to supervise projects for national, European and international science competitions. The Xenoplex STEM Center was awarded with a federal certificate for excellence.
“I wanted to foster an interest in, and a love for science, in elementary, middle-, and high-school students. It is incredibly gratifying to watch our work over the past 14 years bring the excitement of science to so many.” BioEYES Steven is the founder of BioEYES, a non-profit research program that uses zebrafish to excite and educate K-12 students about science and how to think and act like scientists. BioEYES experiments allow students to work as scientists using a student-centered approach. Over the course of 5 days, students collect zebrafish embryos and watch them transform from a single cell to a free-swimming larva with a visibly beating heart and a distinct pigmentation pattern. Students learn about habitats, human and fish anatomy, DNA and cells. BioEYES has reached over 125,000 children worldwide, delivering positive hands-on educational experiences.
“As I developed as a scientist I began to realize the power of the scientific process. I felt emboldened by my growing ability to seek answers to my own questions in a methodical way. I find the most meaning and value in helping others learn to answer their own questions in critical ways.” Journal of Emerging Investigators (JEI) Sarah established the first peer-reviewed publication for middle- and high-school students, the Journal of Emerging Investigators (JEI), which supports and strengthens the next generation of young scientists. JEI offers students the opportunity to focus on communication as an essential part of the scientific process. The peer-review process serves to improve science and strengthen the scientific community, and allowing students early access to this training helps them build their own scientific identity.
“We educate our youth on how to leave their world a better place than they found it. From this, participants practice sustainable habits and are made aware of zero-waste alternatives. This results in campus and household waste reduction, and ultimately cost savings.” Campus Refill Initiative Samantha established the Campus Refill Initiative (CRI), aimed at reducing plastic waste on college campuses by offering sustainable alternatives to single-use plastics. CRI’s mission is to educate the public about plastic waste and alternative to dealing with plastic waste other than recycling. By making small habitual changes that enable avoiding the use of plastic products and replacing them with a longer-lasting and sustainably-sourced materials, society can drastically decrease the demand for plastic product production. By targeting college students, as they begin to form individual habits and make personal lifestyle choices, CRI aims to heighten awareness of the impact of our personal choices on the environment.
“Our method has been adopted by many farmers in Hassan district within 10 years and now it has spread to across India. Reforestation has a significant impact to mitigate the climate change, where plants harvest sun and carbon dioxide and cool the land.” Reforestation to Protect Biodiversity Malali established a method to re-forest any barren or rocky land in a short period, resulting in an increase in vegetation cover, ground-water and biodiversity. Deforestation for agricultural purposes in regions of India have led to water and agrarian crises. In an effort to counteract these effects, 50 acres of barren land were selected to re-plant with native species. Within 5 years, the previously rocky area has become a rainforest and biodiversity has returned, supporting the livelihood of farming families.
“I feel that the history of science offers multitudes of exciting stories that can be told in such a popular medium, and indeed, deserve to be so told.” Das Molekül Bill is the composer of Das Molekül, which tells the story of the discovery of DNA and the race to sequence the human genome through a full-scale work of musical theatre, involving six soloists, a chorus and symphony orchestra. The work focuses on the discovery of the double helix in 1953 and the events that led to this discovery between Francis Crick and James D. Watson in Cambridge, and Maurice Wilkins and Rosalind Franklin in Kings College, London. The second story told by the musical is about the 1998-2000 race between Francis Collins and Craig Venter’s laboratories to sequence the human genome. The sequence of the work is presented in such a way that the two stories are dramaturgically intertwined as a reflection of the double helix itself.
“Accurate science communication is necessary to protect scientific integrity, safeguard research funding and ensure continued innovation to advance society. With our art and research expertise, Squidtoons presents translated information in an approachable yet accurate manner.” Squidtoons Garfield is the creator of Squidtoons, which communicates research through visually appealing, yet scientifically accurate, comics to broaden public understanding of the sciences. Squidtoons distills scientific research into an artful comic, and then enlists expert scientists to edit visual and textual elements in the per-reviewed method. More than 60 comics have been published, covering topics ranging from marine organism anatomy to climate change. Squidtoons comics have been incorporated into an oceanographic college textbook (Essentials of Oceanography, 12th edition, Pearson), a children’s book (Squidtoons: Exploring Ocean Science with Comics, Andrews McMeel Publishing), and an aquarium exhibit (Seymour Marine Discovery Center, Sanata Cruz, USA).
“By connecting artists and scientists for collaboration, we can improve the human condition by accelerating our ability to solve the interdisciplinary problems of our increasingly complex world.” Bridge Art + Science Alliance (BASA) Kyle is the founder of the Bridge Art + Science Alliance (BASA), a program designed to establish collaboration between the art and science departments at USC, creating a synergy between scientific discovery and engaging media. BASA has produced over 20 projects, many premiering at renowned festivals, like SXSW, and winning prestigious awards such as the Association of Medical Illustrators Art & Biology Award.
“This project is important to me because it allows me to give back to a community that has my great respect and appreciation. For others, I hope my sculptures impart creative and scientific inspiration.” Bryan is an Associate Professor and Sculptor whose artwork accurately and artistically represents DNA and protein complexes to inspire public curiosity in science. Bryan’s sculptures are designed using published molecular coordinates (from NCBI) and through discussions with structural biochemists. Artistic elements are presented throughout the sculptures via patinas and various metal work elements. Once finished, these sculptures are donated to a specific researcher or institute in recognition of research achievements and to highlight the beauty and engineering prowess of the natural world.
“Part of the responsibility of science is to combine and use the knowledge and technologies available to solve societal problems, regardless of the scale. Antibiotic resistance fascinates me, because it is a direct and unavoidable result of antibiotic usage.” Determining Antibiotic Susceptibility Nathan developed rapid diagnostics for determining antibiotic susceptibility, which can be used by clinicians at the point of care, before prescribing, to improve outcomes and prevent resistance. This diagnostic is based on precise measurements of the quantity and state of nucleic acids, and has been integrated into microfluidic systems utilizing isothermal amplification to create a 30 minute antibiotic susceptibility test for urinary tract infections. Because this test can be complete in just 30 minutes, it provides clinicians with crucial information they need (and don’t currently have) when prescribing antibiotics.
“Through this work, I hope to inspire a solution that would liberate science in Africa, as well as create the platform that would permit scientists like myself to return home and do good science.” TReND Outreach program&SciComNigera Mahmoud is the founder of the TReND Outreach program and SciComNigera, aimed to enhance public understanding, trust and support for science, and inspiring the next generation of African scientists through outreach activities for school students, teachers, government officials and scientists. The TReND Outreach program has more than 90 science communicators across Africa working to engage students and the public. SciComNigeria was founded to help cennect scientists to the public, promoting local research and science stories, as well as increase the visibility of science in Nigeria.
Using Exonuclease V (RecBCD) to Eliminate Residual Genomic DNA When Purifying Low Copy Plasmids with the Monarch Plasmid Miniprep Kit
Peichung Hsieh, Ph.D., New England Biolabs, Inc.
Introduction The use of low and/or single-copy plasmids to clone large pieces of DNA (up to 200 kb) or to drive expression of slow folding or toxic proteins in E.coli is a commonly used strategy. Purification of low-copy plasmids or bacterial artificial chromosomes (BACs) presents some challenges that are not evident when working with higher copy number plasmids, such as pUC19. The ratio between bacterial gDNA and plasmid DNA is higher, thereby reducing yield of the desired plasmid produced by typical plasmid miniprep protocols. Additionally, elevated levels of host gDNA are often co-purified, reducing the accuracy of quantitation by UV absorbance or dsDNA specific dyes. Neither method can distinguish the contribution from gDNA to the overall nucleic acid content. Co-purification of host gDNA also affects the appearance of the sample when resolving by gel electrophoresis and adds unwanted contaminating DNA for any amplification-based application. Exonuclease V (RecBCD, NEB #M0345) is an exonuclease that degrades both linear ss- and dsDNA, while keeping the circular DNA intact. Treatment of miniprep DNA samples of low copy plasmids with this exonuclease degrades the contaminating gDNA, restoring purity and ease of use in downstream applications.
Results: Three milliliters of an overnight culture of NEB-10 beta competent E. coli cells transformed with pBAC were processed using the Monarch Plasmid DNA Kit and an equivalent Miniprep kit from another vendor. After isolating the DNA, samples were treated with Exonuclease V (RecBCD) and then digested with EcoRI. Samples were run on an agarose gel to assess the quality of the isolated DNA, and whether or not the Exonuclease V-treated DNA could be digested to completion. The Exonuclease V-treated samples showed no gDNA contamination (#3-6) while the untreated samples exhibited a significant amount of gDNA as seen by the smear observed in those samples (#1,2,7,8). These results indicate that Endonuclease V can be used to efficiently degrade contaminating gDNA from plasmid purification steps, including those of low copy number.
Protocol: 1. Transform an endA- strain (e.g. NEB 10-beta, NEB #C3019) with the BAC plasmid DNA and plate outgrowth onto a media plate with appropriate antibiotic. Incubate overnight at 30°C. BACs with CamR require reduced stringency selection. Chloramphenicol levels should be maintained between 10-15 μg/ml on the selective plate. Note: strains with an F’ plasmid are not compatible with BACs or miniF plasmids. 2. Pick a colony, inoculate 10 ml LB + antibiotic, and incubate overnight at 30°C (200-250 RPM). 3. Check OD600 nm (usually it will be around 4 O.D./ml of cells). 4. Harvest 3 ml of the overnight culture and purify the plasmid DNA using the Monarch Plasmid Miniprep Kit (NEB #T1010) following the recommended protocol. 5. In the final elution step, elute the DNA with 30 μl of Monarch DNA Elution Buffer (pre-heated to 50°C). 6. To the eluted DNA, add 4 μl of NEBuffer 4 (10X), 4 μl of 10 mM ATP, and 2 μl of Exonuclease V (RecBCD). Mix reaction and incubate at 37°C for 1 hr. 7. Heat-inactivate the Exonuclease V by incubating at 70°C for 30 min. The plasmid DNA is now ready for restriction enzyme digestion, PCR or transformation. Note: Typically, 30-60 ng of single-copy plasmid can be purified from 3 ml of an overnight E.coli culture with (OD600 = 4 O.D./ml)
Materials • Endonuclease V (RecBCD) (NEB #M0345) • NEB 10-beta Competent E. coli (High Efficiency)(NEB #C3019) • Antibiotic, typically Chloramphenicol • LB Media • Monarch Plasmid Miniprep Kit (NEB #T1010)
pBAC samples exhibit no bacterial gDNA contamination after treatment with Exonuclease V (RecBCD)
Miniprep plasmid DNA samples isolated with the Monarch Plasmid Miniprep kit (N) and a similar kit from a competitor (Q) were either treated (+) or not treated (-) with Exonuclease V, and then digested with EcoRI. The samples treated with Exonuclease V showed no contaminating gDNA and they were correctly cut with EcoRI.
Materials
Protocol
Results
It’s Not Easy Being a Right Whale
Emily Greenhalgh, New England Aquarium
411: The predicted number of right whales remaining in the North Atlantic population.
These behemoths of the sea were once called the “right whales to hunt”, because they swam close to shore, produced high yields of whale oil and baleen, and—thanks to their thick blubber—floated when killed. But now, rather than whalers, the population faces threats from humans in an ever-increasing urbanized ocean. Due primarily to human impacts, the population of these endangered whales has been in decline since 2010. Entanglement in fishing gear, changes in food distribution due to climate change, busy shipping lanes, and ocean noise are just some of the challenges facing the species.
“The species is resilient. We know they can rebound if we just stop killing them.”
Dr. Scott Kraus, Vice President of the New England Aquarium’s Anderson Cabot Center for Ocean Life
“FDR” was photographed in the Bay of Fundy in 2016, entangled in a large amount of fishing gear. He was successfully disentangled by the Campobello Whale Rescue Team. Photographed by Jerry Conway, Campobello Whale Rescue Team. Permit: Collected under Permits issued by Canadian DFO under the Species at Risk Act.
“Caterpillar” was struck by a vessel, which left large propeller cuts on her body. Caterpillar was photographed in the Bay of Fundy on August 5, 2007 by Amy Knowlton, ACCOL, New England Aquarium. Permit: Collected under Permits issued by Canadian DFO under the Species at Risk Act.
This cornerstone has allowed dedicated scientists and researchers to build an array of methods to try to save this species. Since 1999, researchers from the Aquarium’s Anderson Cabot Center have been collecting right whale fecal samples. By examining the levels of glucocorticoid hormones in these samples, Senior Scientist Rosalind Rolland, D.V.M. and her team were able to determine the stress levels of living whales for the first time. From 1999 to 2014, scientists examined samples from 125 different right whales – 113 healthy whales, six chronically entangled in fishing gear, one that was live-stranded, and five killed quickly by vessel strikes. The robust samples allowed Rolland and her team to create a baseline of hormone levels in normal, healthy whales and compare those levels to animals under stress. The researchers found “sky-high hormone levels” in whales entangled in fishing gear. “These levels showed stress from extreme physical trauma,” said Rolland. “It’s an animal welfare issue.” How many of the right whales are undergoing this trauma? A total of 83% of endangered North Atlantic right whales show signs of entanglement, and 59% have been entangled more than once. Entanglements now surpass ship strikes as the main threat to the right whale population. According to Anderson Cabot Center scientists, only a third of severely entangled female whales survive, and those that do survive are less likely to have calves. When in good condition, a right whale can give birth every three years. But with all the threats they are facing now, the birth rate for the population has dropped to an average 10 years between births - births are simply not outpacing deaths. Rolland and other scientists on the Aquarium’s Marine Stress and Ocean Health team built on their pioneering fecal hormone research by studying stress hormone levels in whale blow, baleen, blubber and blood. The data collected from these samples include hormone levels, DNA, marine biotoxins and pathogens, to name just a few—and this helps paint a picture of the overall health of each animal. Knowing how humans are affecting right whale populations is a key factor to protecting them in the future. “We just have to keep from killing them, both directly and indirectly,” said Rolland. Scientists are working with partners across industry, scientific institutions, and the U.S. and Canadian governments to try to save the species from the brink of extinction. From supporting the implementation of ropeless fishing technology to speaking against offshore oil and gas development on our coasts, the right whale community is fighting hard to ensure these iconic whales not just survive, but thrive.
“Because they’re a long-lived species, the right whales can weather short-term events. We have to give them the opportunity.”
Philip Hamilton, Research Scientist
It was humans who nearly destroyed the right whale population, and it is humans who are striving now to save it from extinction. In 1980, New England Aquarium scientists discovered a group of 25 North Atlantic right whales in Canada’s Bay of Fundy. At the time, the species was thought to be almost extinct. In the nearly four decades since that startling discovery, dedicated researchers have been fighting to save this iconic species. For nearly 40 years, members of the Aquarium’s right whale research team have been dedicated to ensuring these whales have the opportunity to survive. Fieldwork, including 39 years of uninterrupted surveys in the Bay of Fundy, has provided invaluable data about right whale behavior, habitat use, and human impacts on the population. The Anderson Cabot Center Right Whale Program also oversees the North Atlantic Right Whale Catalog (rwcatalog.neaq.org), a tremendous collaborative effort of more than 300 individuals and organizations. Right whales are identifiable by callosities, the natural patches on the top of their heads, as well as scars or markings on their bodies. Scientists can recognize individual whales from these marks in thousands of photographs, connecting important information about the population, such as location, mortality, health, and reproductive success. The Right Whale Catalog, which is linked with human impact studies, visual health assessments, and genetic and hormone analysis, is the cornerstone of right whale research. Genetic samples from biopsies have also helped scientists estimate the original size of the right whale population before commercial hunting, and even shed light on how few calving-age females there were during the population’s lowest point. About 80% of the right whale population has been sampled, and between the genetic database and the Right Whale Catalog, scientists know entire family histories for many of these whales.
